Blue-Light Sensing in Arabidopsis: In Vivo Microscopy
of Subcellular Localization, Endocytosis, and Vesicular
Recycling of the Blue-Light Receptor Phototropin 1
Dissertation
zur
Erlangung des Doktorgrades (Dr. rer. nat.)
der
Mathematisch-Naturwissenschaftlichen Fakultät
der
Rheinischen Friedrich-Wilhelms-Universität Bonn
vorgelegt von
Yinglang Wan
aus
Sichuan, China
Bonn 2008
Angefertigt mit Genehmigung der Mathematisch-Naturwissenschaftlichen
Fakultät der Rheinischen Friedrich-Wilhelms-Universität Bonn
1. Referent: Prof. Dr. František Baluška
2. Referent: Prof. Dr. Diedrik Menzel
Tag der Promotion: 13.02.2009
Erscheinungsjahr: 2009
2
Table of Contents
List of Figures
List of Tables
Abbreviations
1.
Introduction
1.1
Visible Light and Plant Photoreceptors
1
1.2
Phototropism and Polar Auxin Transport
3
1.3
Phototropism and Interaction between Gravi- and Phototropism at
Root Tip
7
1.4
Phototropins
9
1.5
Modulation of PIN-formed Protein Localization
13
1.6
Introduction to the Chemicals Used in this Study
15
1.6.1 Brefeldin A
15
1.6.2 Cycloheximide
16
1.6.3 Latrunculin B
16
1.6.4 Wortmannin
16
1.6.5 FM4-64 (Synaptored) and FM1-43
16
2.
Materials and Methods
2.1
Plants Materials
18
2.1.1 Growth Condition
18
2.1.2 Genetically Transformed Arabidopsis Lines
18
2.2
Chemicals
19
2.3
Methods
20
2.3.1 Crossing of transgenic lines of Arabidopsis
20
2.3.2 Methods to Produce Transgenic Line of Arabidopsis
20
3
2.3.3 Stepwise Staining of Membrane Fractions.
22
2.3.4 Confocal Laser Scanning Microscopy and Blue-Light illumination
23
2.3.5 Analysis of Whole Seedling Fluorescence
23
2.3.6 Histochemical ß-Glucoronidase (GUS) staining
24
2.3.7 Determine the New Protein Synthesis by
35
S-Methionine
2.3.8 Phototropic Analysis
3
Results
3.1
Subcellular Localization Phototropin 1-GFP in Etiolated Seedlings of
Arabidopsis thaliana
24
25
27
3.1.1 Phototropic Analysis of proPHOT1::PHOT1::GFP Tansformed
Seedlings
27
3.1.2 Cellular and Subcellular Localization of PHOT1::GFP Distribution in
Etiolated Seedlings
28
3.1.3 Distribution in the Cotyledons
29
3.1.4 Distribution in the Hook
32
3.1.5 Changes from Hook to Transition Region as Visualized in
Cross-Section
34
3.1.6 Distribution in Elongating and Mature Hypocotyl Tissues
35
3.1.7 Distribution in the Shoot–Root Transition Zone
36
3.1.8 Distribution in Mature and Elongating Root Tissues
37
3.1.9 Subcellular Distribution of PHOT1::GFP in the Root Apex
37
3.2
40
BL-Induced Vesicular Re-localization of PHOT1::GFP Molecules
3.2.1 Hypocotyl cell
40
3.2.2 Cotyledon Cells
44
3.2.3 Root Cortical Cells
45
3.2.4 BL-Induced Movement and Dark Recovery in the Presence of a
Protein Synthesis Inhibitor
46
4
3.2.5 BL-induced Internalization of Membrane-associated PHOT1 via
Endocytosis
48
3.2.6 Endosomal PHOT1 Undergoes BL-Sensitive Vesicular Recycling
3.3
49
The Endosomal Recycling of PIN-formed Proteins is Sensitive to the
BL Illumination
53
3.3.1 PIN2 is Essential for the BL-Induced Phototropic Responses in Roots 53
3.3.2 Brefeldin A-Inhibited BL Induced Responses in Arabidopsis
54
3.3.3 BFA-Sensitive Recycling of PIN2 is Dependent on BL Illumination
55
3.3.4 BL Influences the Localization of PIN 1 and PIN2 Protein
57
3.3.5 Endocytosis and Fusion of Endosomal Vesicles are Accelerated by
BL Illumination
60
3.3.6 NPH3::GFP has Polar Localization at Root Tip Region
62
3.3.7 BL Illumination Changes Size of Meristem and Transition Zone
63
3.3.8 Asymmetric Auxin Distribution are Affected by Lateral BL Signals
64
3.3.9 PIN3 is an Essential Factor to Root Phototropism
65
4
Discussion
4.1
The
tissue
distribution
pattern
of
PHOT1
correlates
with
Physiological Responses
4.1.1 Expression
of
proPHOT1::PHOT1::GFP
66
Gene
Rescues
the
Phenotype of phot1-5 Mutant
4.1.2 Subcellular Distribution of PHOT1
67
69
4.1.3 Cross-Wall Localization of PHOT1 in Root Transition Zone
Suggests Roles of PHOT1 in Polar Auxin Transport
4.2
71
BL-Induced Relocalization of PHOT1::GFP is achieved by Receptor
Mediated Endocytosis
73
4.2.1 Blue-Light Effects on PHOT1::GFP Subcellular Distribution
73
4.2.2 Internalization of PHOT1 is Accomplished via Endocytosis
74
5
4.2.3 ARF1-GTPase, the Key Regulator of Endocytosis, is Modified by
PHOT1
75
4.2.4 Endocytosis of PHOT1::GFP is Accelerated by BL Illumination
76
4.2.5 Signalling Endosomes as Integrators of Environmental Signals?
77
4.2.6 Model to Describe the PHOT1-Mediated Signalling by Endosomal
Vesicular Relocalization
4.3
78
Intracellular Localization and Endosomal Recycling of PIN-formed
Proteins are Determined by BL Illumiation
80
4.3.1 Light Signal Changes Subcellular Localization of PIN1 and PIN2
Proteins
80
4.3.2 ARF Emerges as a Key Factor in the BL Induced Endocytic
Trafficking
81
4.3.3 Root Phototropism
83
5.
86
Outlook
Summary
87
References
88
Acknowledgement
102
Curriculum Vitae
6
List of Figures
Figure 1.
Schematic description of endosomal recycling of PIN proteins.
4
Figure 2.
Vesicle transport of auxin mediated by PIN proteins.
5
Figure 3.
The polar auxin transport under gravitropic stimulation.
6
Figure 4.
Schematic depiction of growth zone in root tip of Arabidopsis
thaliana.
7
Figure 5.
Schematic figure shows the domains of PHOT1 protein.
9
Figure 6.
Proposed photocycle of LOV domains from PHOTs.
10
Figure 7.
Schematic model describes the activation of C-terminal kinase
domain of PHOT2 molecule.
Figure 8.
11
A model to describe the signal transduction via PHOT1/NPH3
pathway.
Figure 9.
12
PINOID affects the development of inflorescence and polar
localization of PIN proteins.
14
Figure 10. PHOT1::GFP expression in dark-grown Arabidopsis seedlings. 28
Figure 11. Single
confocal
optical
section
showing
PHOT1::GFP
localization in the Cotyledon and apical hook region.
30
Figure 12. Distribution of PHOT1::GFP in the cotyledons of 4days old
seedlings.
31
Figure 13. Localization of PHOT1 on surface of chloroplasts in mesophyll
cells.
Figure 14. Localization of PHOT1::GFP in the apical hook region.
32
33
Figure 15. Computer-reconstructed cross-sectional images of hypocotyl
cortex and epidermis.
34
Figure 16. Localization of PHOT1::GFP in the elongation zone.
35
Figure 17. Localization of PHOT1::GFP at the shoot–root transition zone.
36
Figure 18. Distribution of PHOT1::GFP in roots of 4 days old etiolated
seedlings.
38
Figure 19. Blue-light-induced changes in PHOT1::GFP distribution in
cortical cells.
41
7
Figure 20. Sensitivity of PHOT1::GFP re-localization in hypocotyl cortical
cells to blue light.
42
Figure 21. Disappearance of PHOT1::GFP from cytoplasm of hypocotyl
cortical cells with time in the dark.
43
Figure 22. Blue-Light-Induced re-localization of PHOT1::GFP in hypocotyl
cortical cells is sensitive to total photon fluence.
43
Figure 23. Blue-light-induced re-localization of PHOT1::GFP in cotyledon
epidermal and mesophyll cells.
44
Figure 24. Blue-light-induced PHOT1::GFP re-localization in root cortical
cells.
46
Figure 25. Effect of cycloheximide on blue light-induced PHOT1::GFP
re-localization and subsequent recovery in darkness.
47
Figure 26. The Released PHOT1::GFP signals from PM of shoot
epidermal cells is colocalized with endosome tracer FM4-64.
48
Figure 27. The Released PHOT1::GFP signals from the PM of root
cortical cells co-localize with endosomes.
Figure 28. Colocalization
of
PHOT1::GFP
and
49
FM4-64
within
BFA-induced endosomal compartments.
51
Figure 29. Still images from a time series of cells observed during BFA
treatment.
Figure 30. BFA-sensibility of root cortical cells in different conditions.
51
52
Figure 31. PIN2 is essential for root phototropic responses but has no
effects in hypocotyl.
Figure 32. Brefeldin A inhibits responses to light and gravity.
53
54
Figure 33. BFA-sensitive recycling of PIN2 is depended on blue light
illumination.
56
Figure 34. PIN1::GFP Localization and Relocalization in Root Cells under
Light and Dark signals.
58
Figure 35. PIN2::GFP localization and relocalization in root cells under
light and dark conditions.
59
Figure 36. Sequential membrane labelling reveals an effect of light on the
8
fusion of early and late endosome under BFA treatment.
61
Figure 37. Cellular localization of NPH3::GFP in the root tip region.
62
Figure 38. Expression pattern of proCYC B1::GUS in the root tip region.
63
Figure 39: Expression pattern of proDR5::GFP at the root tip.
64
Figure 40. Analysis of phototropism in pin3-3 mutants.
65
Figure 41. Model to explain PHOT1 movement under blue light
illumination.
Figure 42
79
Model to explain how blue light modifies PIN-mediated auxin
transport.
82
List of Tables
Table 1
List of chemicals
Table 2.
Complementation of phototroism in etiolated Arabidopsis
hypocotyls by PHOT1::GFP.
Table 3.
Table 5.
27
Expression level of PHOT1::GFP in various tissues of etiolated
Arabidopsis seedlings.
Table 4.
19
29
Speed of BFA induced compartment formation in different cell
types.
50
Brefeldin A inhibits responses to light and gravity.
55
9
Abbreviations
AGC
Protein kinase A, cyclic GMP-dependent protein kinases
and protein kinase C
APS
Ammonium persulfat solution
ARF
ADP-Ribosylation Factor
ARF-GEFs
(ARF-Guanine-nucleotide Exchange Factors)
BFA
Brefeldin A
BL
Blue Light
CEZ
Central Elongation Zone
CHX
Cycloheximide
Col 0
Columbia 0
CRY
Cryptochrom
DEZ
Distal Elongation Zone
DMSO
Dimethyl Sulfoxide
EDTA
Ethylenediaminetetraacetic acid
F-actin
Actin Filaments
Fig.
Figure.
FM1-43
N-(3-triethylammoniumpropyl)-4-(4-(diethylamino)
styryl)pyridinium dibromide
FM4-64
N-(3-triethylammoniumpropyl)-4-(6-(4-(diethylamino)
phenyl) hexatrienyl)pyridinium dibromide
GL
Green Light
GFP
Green fluorescence protein
GUS
ß-Glucoronidase
IAA
Indole-3-acetic acid
LB
Latrunculin B
LOV
Voltage, Oxygen and Light
MS
Murashige and Skoog
NPA
N-naphthylphthalmic acid
NPH
Nonphotptropic hypocotyls
10
PAGE
Polyacrylamide gel electrophoresis
PAT
Polar auxin transport
PHOT
Phototropin
PHY
Phytochrom
PID
PINOID
PINs
PIN-formed proteins
PM
Plasma membrane
PVC
Prevascular Vesicles
RC
Root Cap
RME
Receptor-Mediated Endocytosis
RL
Red Light
RTZ
Root Transition Zone
SDS
Sodium dodecyl sulfate
TEMED
N, N, N', N'-tetramethylethylenediamine
TGN
Trans Golgi Network
TIBA
2,3,5-triiodobenzoic acid
Tris
Tris (hydroxy methyl) aminomethane
WS
Wassilewskija
WT
Wild-Type
11
1.
Introduction
1.1
Visible Light and Plant Photoreceptors
Light exists universally. As Albert Einstein, Luis de Broglie and many other
scientists discovered in the 20th century, light has a wave-particle duality. The energy
of light is carried by the elementary particle named photon, which is an undividable
entity and therefore it is also called quantum. Einstein presented the following
formula to describe the relationship between the photon-carried light energy (E) and
the frequency (f) of electromagnetic light wave (Formula 1). Luis de Broglie
formulated the hypothesis that not only light, but all matter, has the wave-particle
duality. He described the relation between wavelength (λ) and momentum (ρ) as in
Formula 2. These two formulas are equal to each other and can be linked by the light
speed in vacuum (c) as Formula 3 and Formula 4 (reviewed by Bennett, 2005).
E
=
hf
(Formula 1)
λ
=
h/ρ
(Formula 2)
ρ
=
E/c
(Formula 3)
λ
=
c/ f
(Formula 4)
(Planck’s constant, h= 6.626 × 10-34 J s, light speed, c= 299,792,458 m/s)
Light and the majority other kinds of electromagnetic waves on the earth surface
are radiated from the nearest star, the sun. It affects all aspects of life on our planet.
During billions of years of evolution, life forms have developed several molecular
systems to take advantage of sunshine directly and indirectly. Among them, the
advent of oxygenic photosynthesis is one of the most important events in evolution of
life on earth (Xiong et al., 2000). About 3.5 billions years ago, ancient cyanobacteria
invented a mechanism to transform light energy into chemical energy using water as
the electron donor (Xiong et al., 2000, Dyall et al., 2004). About 1.2 billions of years
ago, such ancient cyanobacteria were engulfed by unicellular eukaryotes, giving rise
to a stable endosymbiosis, and eventually the endosymbionts transformed in to
chloroplasts (Des Marais, 2000, Xiong et al., 2000). Due to chloroplasts, light became
the primary energy source for algae and the green plants and all the animals and
heterotrophs, which live on green plants directly or indirectly, depend on this energy
source as well.
1
Plants need to orient themselves in space and time in order to perceive light
efficiently. Therefore, light does not only serve as energy source, but also as an
environmental signal for plant lives. Light signals from the environment can influence
the direction of growth in organs such as stems and roots, bending towards or away
from the light source. These responses are called phototropism.
In Arabidopsis, the shoots react positive and the roots negative to incident blue
light (BL) signals (reviewed by Iino, 2006), while red light (RL) causes reverse
responses (Molas and Kiss, 2008, Kiss et al., 2003). Both qualities of light may also
induce photomorphogenesis. Furthermore, excess light is destructive. Leaves may
fold or rotate relative to the light vector, chloroplasts within the cells may reorient and
tissues may accumulate photoprotective substances in order to prevent photo
damages (Koller, 2000). All these implicate, that plants must have systems to sense
and analyze light signals.
But unlike animals, plants do not have sensory organs such as eyes. In fact, the
term “light” is derived from the physiological mechanism of the human eyes, which
can detect electromagnetic waves with the wavelength range between 380-800 nm.
Therefore this part of the electromagnetic wave spectrum is becoming visible light, or
simply light. There are three kinds of cone cells in the human eye that are sensitive to
three different ranges of wavelength (reviewed by Solomon and Lennie, 2007), and
hence light is divided into three different color groups: blue light (BL, 380-500 nm),
green light (GL, 500-600 nm) and red light (RL, 600-800nm). Plants detect a similar
spectrum of lights as humans do, but mostly blue and red light signals, while they are
blind for green light (Folta and Maruhnich, 2007).
Three families of photoreceptors have been discovered in plants and they are
characterized in greater detail in the model plant Arabidopsis: phototropins (PHOT),
cryptochromes (CRY) and phytochromes (PHY) (reviewed by Lariguet and Dunand,
2002). Phototropins and cryptochromes are activated by blue and ultraviolet-A (UV-A)
light signals, while phytochromes are sensitive to red and far-red lights. The
phototropin family has two members, named PHOT1 and PHOT2, which mediate
most BL-initiated responses, such as phototropism, stomata opening, leaf expansion
and chloroplast movement (reviewed by Christie, 2007). Cryptochrome has two
known members, named CRY1 and CRY2. They regulate organ movements,
physiological and morphogenetic responses in plants, such as inhibition of hypocotyl
elongation, anthocyanin accumulation, promotion of stem and internode elongation,
2
circadian clock and day length perception, the latter of which helps plants to optimize
their flowering time (reviewed by Li and Yang, 2007). The phytochome family has 5
members, named PHYA, PHYB, PHYC, PHYD and PHYE (Sharrock and Quail,
1989). They regulate several aspects of plant development and growth, such as seed
germination, seedling de-etiolation, neighbour perception and avoidance, and flowing
(reviewed by Schepens et al., 2004).
In this thesis work, my studies are focused on the BL receptor PHOT1, and
PHOT1-mediated phototropic responses.
1.2
Phototropism and Polar Auxin Transport
Phototropism is the bending response of plant stem or root tissues under unilateral
light illumination. The observation of this phenomenon is traced to the ancient
civilizations, but scientific studies have only been started at the beginning of the 19th
century (Poggioli, 1817, thanks to Prof. Winslow Briggs to unearth this history and
introduced to me). Since then, numerous studies have been dedicated to the
mechanisms of phototropic response. The most widely accepted theory was
presented by Cholodny and Went independently (Went, 1928, Cholodny, 1926). They
suggested that the asymmetric redistribution of the growth hormone auxin, namely
indole-3-acetic acid (IAA), in response to gravity or unilateral light lead to organ
curvature by causing unequal rates of cell elongation at opposite sides of the organ.
After decades of physiological studies, this theory has recently been supported more
directly by genetic and molecular biological studies addressing the polar auxin
transport (PAT) as a central aspect of photo- and gravitropism. Molecular
components have been discovered such as auxin efflux and influx transporters and
transport facilitators in plant cells, and their roles in the tropic responses have been
characterized (Friml et al., 2002, Blilou et al., 2005). Members of the PIN protein
family have been identified as auxin efflux facilitators in Arabidopsis and their
intracellular localization at the apical and/or basal plasma membrane (PM) domains
in the cell files has been determined. PIN1 cycles between the PM domain and the
endosomal compartment (Fig. 1) (Bonifacino and Jackson, 2008). Evidence for this
cycling behavior comes from work with inhibitors of auxin transport. For example, Nnaphthylphthalmic acid (NPA) and 2,3,5-triiodobenzoic acid (TIBA), as inhibitors of
cellular auxin export, and brefeldin A (BFA) (inhibitor of protein secretion). All three
3
block auxin transport by inhibiting the endocytic recycling of PIN proteins (Geldner et
al., 2001, 2003).
Brefeldin A is an antibiotic produced by the fungal organism Eupenicillium
brefeldianum (Fujiwara et al., 1988). BFA binds to ARF-GEFs (ADP-Ribosylation
Factor-Guanine-Nucleotide Exchange Factors) specifically, therefore blocks the
activation of ARF by stopping the phosphate exchange between ARF-GDP (inactive)
and ARF-GTP (active). ARF activation by GEF is important at least in two situations:
the first one is when COPII vesicles form to be exported to the cis-Golgi cisterna.
Inhibition of this GEF by BFA leads to inhibition of secretion; the second situation
involving GEF-factors is when vesicles bud off the endosome to be exported to the
plasmamembrane. This latter member of the ARF-GEF factor family in plants is
named GNOM. GNOM is associated with endosome membranes and the
Figure 1. Schematic description of endosomal recycling of PIN proteins (Bonifacino
and Jackson, 2003).
BFA inhibits the GNOM-mediated activation of ARF. ARF plays critical roles in the protein
transportation via vesicle pathways. COP1, GGA1 and other coat proteins are involved. The
recycling of PIN proteins is suggested to be related to this process.
4
Figure 2. Vesicle transport of auxin mediated by PIN proteins (Baluška et al., 2003).
Detailed view of postulated PIN1 recycling at the apical pole of a hypothetical root cell. PIN1
(yellow circles) recycles between the apical plasma membrane (PM) and endosome (E). In (a),
the conventional concept is shown, with PIN1 performing auxin (black dots) export (indicated with
arrows) only when inserted into the plasma membrane. This model does not explain the transient
nature of the PIN1 localization to the plasma membrane. In (b), the newly emerging model is
summarized, based on the current breakthrough papers as well as on older data. According to this
concept, PIN1 is the vesicular transporter that loads auxin (black dots) from the cytoplasm into
vesicles and endosomes, while localization of PIN1 to the plasma membrane is only transient
owing to its vesicular secretion and rapid retrieval back into internal membranes. In (c), the
synaptic vesicle is shown in comparison with a putative auxin-accumulating vesicle. Both types of
vesicles accumulate either glutamate or auxin, using a proton gradient as the driving force for
crossing their limiting membranes.
blocking of GNOM by BFA results in disrupted membrane recycling between
endosomes and plasma membranes whereas endocytosis from the plasma
membrane is not blocked. Block of secretion and of endosomal recycling lead to a
block of endosomal vesicular trafficking and consequently to the formation of BFAinduced compartments (Anders et al., 2008, Teh and Moore, 2002). Since PINproteins recycle between plasma membrane (PM) and endosomal compartment, their
5
correct distribution in the cell is also blocked and therefore processes depending on
PIN-function are also blocked by BFA (Fig. 1, Bonifacino and Jackson 2008, Inoue
and Randazzo, 2007, Dhonukshe et al., 2007). Therefore Baluška et al. (2003)
hypothesized that auxin is transported in a vesicle manner inside plant cells (Fig. 2).
This hypothesis about adjustable auxin transport in endosomal vesicles are
supported by current studies of our group (Schlicht et al., 2006).
PIN-formed proteins (PINs) are relocalized under changes of both gravity and
light conditions. For example, a member of the PIN family - PIN3 – has been reported
to play an important role in the redistribution of auxin in the root cap following
gravistimulation (Friml et al., 2002). After reoriention of the roots from vertical to
horizontal position, the PIN3 proteins relocated from the horizontal to the vertical PMs
Figure 3. The polar auxin transport under gravitropic stimulation (Abas et al., 2006).
A: Auxin flow in the vertical position (blue arrows). PIN1 proteins are localized at the cross-walls of
central cylinder cells, driving the apical transport of auxin. PIN2 proteins are at the polar domains
of the plasma membrane in cortical and epidermal cells, and they transport auxin basally. PIN3
are localized in the root cap statocytes where they act as the re-distributor of auxin.
B: Auxin flow in the horizon position, changed by the relocalization of PIN3 proteins (green). Most
of auxin is flowing to the lower part, with help from PIN3 proteins.
C: The auxin flow in the lower part of the elongation zone. The cellular concentration in the cells in
this side is much higher than at the upper side. Cell elongation is inhibited and results in the
root apex bending response to direction of gravity.
6
of the columella cells. This observation suggests that the PIN3 proteins determine the
auxin distribution under reorientation of the organ. Another member of the PIN family,
PIN1, becomes re-located in cells on the shaded side of the hypocotyl of wild-type
Arabidopsis but is not similarly re-localized in a phot1 mutant (Blakeslee et al., 2004).
The localization of PIN2 is affected by light conditions as well. PIN2::GFP signals are
stimulated inside vacuole in dark-grown seedlings while the seedlings grown under
BL illumination did not have such pattern of localization (Laxmi et al., 2008). The
mechanisms and the functional roles of this PIN2 relocalization are still unclear. And
the roles of PINs in phototropic responses are still unknown either. Recently, the
network built up by PIN1, PIN2 and PIN3 proteins and their roles in polar auxin
transport are gravitropic stimulation is described in the Fig. 3 (Abas et al., 2006).
1.3
Phototropism and Interaction between Gravi- and Phototropism at Root
Tip
Though the functions of PINs in Arabidopsis root under gravitropic stimulation are
relative well studied, the phototropism of plant roots are only poorly studied. Because
roots generally grow beneath the soil and light can only penetrate several millimeters
into the soil, root phototropism is usually of little importance. However, when grown in
light, root phototropism is generally masked by gravitropism. Although an early study
on root phototropism surveyed 152 species and found 50% of these plants have
negative phototropism in roots (Hubert and Funke, 1937), only a few of them have
been characterized by modern technique in the absence of gravity.
Figure 4. Schematic depiction of growth zone in root tip of Arabidopsis thaliana.
Regions are colored in different colors.
White: root cap; blue: quiescent center; red: meristem; yellow: root transition zone (distal
elongation zone), green: central elongation zone.
7
Elongation rate of different tissues in phototropic response was detailed (Orbovik
et al., 1993). A computer-feed back system was applied to observe the phototropism
and gravitropism responses of Arabidopsis roots (Mullen et al., 2000). The response
to light occurs at the central elongation zone (CEZ), while the roots respond to gravity
at the root transition zone (RTZ), also known as distal elongation zone (DEZ)
(distribution of growth zones is shown in the Fig. 4). The time that the roots need to
respond is different too: the response to light is slower (40 minutes) than the
response to gravity (10 minutes). These results suggest that the phototropic and
gravitropic responses may share pathways of signal transduction, but each may
evoke specific pathways as well, though it is considered, that the root cap is the
organ for sensing both of gravity and light. Mullen and co-workers used optical fiber
to illuminate the root cap and the RTZ of Zea mays (Mullen et al., 2002). They found
that the illumination of the root cap (RC) led to significant, negative phototropic
bending, but that the illumination to the RTZ did not. They concluded that the
perception of light and the response to it are spatially separated.
The relationship between gravitropism and phototropism in a starchless mutant of
Arabidopsis has been observed and analyzed (Vitha et al., 2000). It has been
discussed, that the phototropism in the root region is a complex process determined
by the intensity, direction and rhythm of illumination. Not surprisingly, the
phototropism is unmasked by removing the gravitropic responses. Phototropic
responses in starchless mutants are three times more intense than in the wild type
root apices. Furthermore, roles of different light receptors in root tropic responses
have been discovered by modern molecular biology. Roots of Arabidopsis are
bending away from BL, but towards RL. This positive response to RL is mediated by
phytochrome A and B (Kiss et al., 2003). Lariguet and Fankhauser have analyzed the
interaction between gravitropic responses by using photoreceptor mutant plants. The
Null mutant phot1-5 shows non-gravitropic phenomena under weak BL illumination,
while phot2 and phot1phyA mutants have normal gravitropism under the same
conditions. They suggest that activation of PHYA under BL inhibits gravitropic
responses (Lariguet and Fankhauser, 2004), and that PhyA acts as a positive
regulator of BL-caused phototropism, and thus it influences the phototropism induced
by the PHOT1 BL receptor. It is therefore safe to conclude, that the signaling
pathways under BL and RL interact with each other and that the gravitropic and
phototropic signals influence each other, too (Correll et al., 2002, Folta et al., 2001,
8
Kiss et al., 2001). PKS1 (Phytochrome kinase substrate 1) links to PHOT1 and is
supposed to be one of the interaction points in the complex signal transduction and
response pathways (Schepend et al., 2008, Lariguet et al., 2006).
1.4
Phototropins
How can plants sense the direction of light? A gradient of light intensity between
the illuminated and the shaded sides of a plant would be a logical answer to this
question. Optical fibers have been used to measure the light gradient in the
coleoptiles and hypocotyls of seedlings irradiated by unilateral BL (Vogelmann and
Haupt, 1985). It has been determined, that light reaching the shaded side is 4-8 times
weaker than the light hitting the illuminated side of maize hypocotyls. In order to
sense the direction of light, plant cells must have mechanisms to sense and analyze
this light gradient created by shading effects.
In the 1990s’, Briggs and his colleagues have found that the level of membrane
protein phosphorylation showed a gradient between illuminated and shaded side
(Short et al., 1994, Palme et al., 1993), and therefore these results provided the
evidences pointing at the involvement of a protein kinase . In 1995, mutant alleles
named nph (nonphotptropic hypocotyls) were presented (Liscum and Briggs, 1995
and 1996). Among them, the NPH1 gene encodes a flavoprotein with putative light
and redox sensing domains, the LOV domains, which are activated by voltage,
oxygen and light signals (Fig. 5). PHOT1 is phosphorylated even under weak BL
illumination (mechanism is shown in the Fig. 6). Later, researchers found that the
gene on the NPH2 locus encoded a NPH1-like protein (NPL1), which was activated
and phosphorylated under stronger light illumination (Sakai et al., 2001, Kagawa et
al., 2001). Therefore, NPH1 and NPL1 were assigned to the same protein family and
renamed phototropin 1 and 2 (Briggs et al., 2001).
Figure 5. Schematic figure shows the domains of PHOT1 protein.
PHOT1 has two LOV domains at N-terminus, each LOV domain binds a FMN molecule noncovalently. There is a ser/thr kinase domain at C-terminus.
9
Figure 6. Proposed photocycle of
LOV domains from PHOTs
(Matsuoka et al., 2007).
Absorption of blue light results in the
generation of the singlet excited-state
FMN. Intersystem crossing results in
the formation of the triplet-state, and
this excited state is stabilized by
protonation of the N5 atom of the
isoalloxazine (high lighted in red) ring
through the abstraction of a proton from
the conserved Cys in the LOV domain
polypeptide. This protonation of the
triplet
state
FMN
increases
the
electrophilicity of the C(4a) atom and
promotes nucleophilic attack of the thiol
anion at this position, resulting in the
formation of the cysteinyl-FMN adduct
(Arrows right side). This adduct is
completely reversible in darkness. The
catalytically reactive N5 and C(4a)
atoms are highlighted on the ground
state
molecule,
and
their
excited
orbitals are highlighted in blue on the
singlet and triplet state molecules.
(arrows left side).
Phototropins are the most essential photoreceptors for the phototropic signaling.
They trigger the BL-mediated signal transduction pathways. Both of them are
sensitive to BL signals with peak wavelength at 450 nm. Already very weak BL
(>0.01 µmol·m-2·s-1) can activate PHOT1, whereas the PHOT2 has less sensitivity to
BL and requires much higher doses (> 1µmol·m-2·s-1) (Sakai et al., 2001). Besides
the phototropic response, phototropins mediate also other BL mediated responses in
plants. For instance, PHOT2 mediates dispersal of chloroplasts under strong BL,
while both PHOTs together mediate intracellular chloroplast movements under weak
BL (Kagawa et al., 2001, Suetsugu et al., 2005). Moreover, both PHOTs mediate
stomata opening/closing (Kinoshita et al., 2001, 2003), expanding of leaves
(Sakamoto and Briggs, 2002) and inhibition of hypocotyl elongation (Christie, 2007).
10
Figure 7. Schematic model describes the
activation of C-terminus kinase domain of
PHOT2 molecule (Matsuoka et al., 2007).
Blue light releases of the FMN from LOV2
domain, which inhibits kinase activation at
ground status in dark condition. After blue
light illumination, C-terminus kinase is freed
from the LOV2. Sites of phosphorylation of
ser/thr amino acids are at N-terminus.
Actually, 8 sites have been discovered
(Salomon et al., 2003). Except phototropin
itself, still no other substrate is known.
According to the photochemistry studies and structure analysis of PHOTs, the BL
signals are received by the two N-terminus LOV domains, which then activate the
ser/thr kinase domain at the C-terminus (Freddolino et al., 2006, Eitoku et al., 2007).
LOV proteins exist widely in all kinds of eukaryotic organisms. Among all the LOV
proteins, PHOTs are the only two members with two LOV domains for some
unknown reasons. It is suggested that the LOV2 domain plays an essential role in the
activation of the C-terminus kinase (Matsuoka et al., 2007), while the LOV1 domain
helps in the dimeration of PHOT molecules (Salomon et al., 2004). Furthermore, the
only known in vivo substrate of C-terminus ser/thr kinases is phototropin itself, though
a common ser/thr kinase substrate, casein, is phosphorylated by the kinase domain
of PHOT2 in vitro (Matsuoka and Tokutomi, 2005). Therefore, the signals are not
transduced simply by the kinase cascade pathways. Though the exact mechanisms
of phototropins-mediated pathways are still unknown, several cooperation partners of
PHOTs were proved to mediate different BL responses. For example, NPH3, RPT2
(root phototropism 2) and CPT1 (coleoptile phototropism 1) belong to the same
protein family and interact with PHOT1 directly (Motchoulski and Liscum, 1999, Inada
et al., 2004, Haga et al., 2005). It has been suggested, that NHP3/RPT2/CPT1 act as
scaffold proteins and mediate the signal transduction in phototropic responses, RPT2
can also control stomata movements (Inada et al., 2004). But neither of them plays
roles in chloroplast movement responses. The mechanisms of the pathway via
PHOT1/NPH3 have been described relative clearly (Motchoulski and Liscum, 1999,
Pedmale and Liscum, 2007). Yeast two-hybrid and in vitro immunoprecipitation
11
Figure 8. A model to describe
the signal transduction via
PHOT1/NPH3 pathway.
(Pedmale and Liscum, 2007).
NPH3 binds to PHOT1 directly
via the coil-coiled domain (CC).
LOV2 in the dark prevents the
activation of the C-terminus kinase
domain. Blue light illumination
causes activation of the kinase
domain of PHOT1 which is
autophosphorylated. Blue light
mediates dephosphorylation of
NPH3 proteins via the regulatory
subunit of PPase. The BTB domain
at the N-terminus of NPH3 is
activated by dephosphorylation.
This then leads to downstream
signal transduction.
...
suggests that the coiled-coil domain at C-terminus of NPH3 interacts with the Nterminus of PHOT1. BL irradiation causes the dephosphorylation of NPH3 via
PHOT1-dependent pathways. They presented a common model to describe the
PHOT1 mediated signal transduction via the PHOT1/NPH complex (Fig. 8).
Ion channels (Ca2+, K+, Na+ , Cl–, and H+) are involved in the PHOT-mediated
responses as well (Babourina et al., 2003; Baum et al., 1999; Fuchs et al., 2003;
Harada et al., 2003; Stoelzle et al., 2003). PHOTs seem also to affect the
distributions of myosin motor protein on the chloroplast surfaces (Krzeszowiec et al.,
2007).
Recent clues in the discovery of the PHOTs mediated pathways are coming from
the analysis of localization of PHOTs. Both PHOTs do not have transmembrane
domains, but early studies with immuno-blotting methods indicated that the PHOTs
are co-localized in the membrane fractions in etiolated Arabidopsis plants (Liscum
and Briggs, 1995). Sakamoto (2002) observed and described the cellular localization
12
of PHOT1 proteins with PHOT1::GFP reporter constructs stably expressed in
Arabidopsis. Interestingly, the illumination with the blue laser in the confocal
microscope caused the release of PHOT1::GFP signals from plasma membrane (PM)
into the cytoplasm. Knieb and colleagues reported that about 20% of the whole
PHOT1::GFP was released from PM into the cytoplasm after illumination (2004), the
level and speed of this process is dependent on the intensity and time of BL
illumination (Wan et al., 2008). PHOT2 shows a similar behavior of BL induced
internalization from the PM into cytoplasm and redistribution to the Golgi apparatus
(Kong et al., 2005).
1.5
Modulation of PIN Localization
Polar localization and endosomal recycling of PIN proteins between PM and
endosomal vesicles provide a plausible model to explain the mechanism for PAT in
plant cells (Baluška et al., 2003, Blilou et al., 2005). The PINOID (PID) protein, a
Ser/Thr kinase (Christensen et al., 2000, Friml et al., 2004) as well as the PP2A
phosphatase (Michniewicz et al., 2007) act as modulators of PINs by controlling their
phosphorylation and dephosphorylation. PID regulates the development of organs by
enhancing polar auxin transport (Benjamins et al., 2001). For example, the
development of inflorescences and lateral roots are affected by the PID kinase
(Christensen et al., 2000, Benjamins et al., 2001). PID also affects the polarity of PIN
localization (Friml et al., 2004). In shoot cells of wild-type plants (WT), PIN1 is
localized on the apical membrane, whereas in the pinoid mutant plants PIN1 have
basal membrane localization. In the root, where the PID gene is not expressed under
normal conditions, PIN1, 2 and 4 proteins are basally localized. However, ectopic
expression of PID in root cells results in apical membrane localization of PIN2 in root
cells (Friml et al., 2004, reviewed by Kaplinsky and Barton, 2004, Fig. 9). There is still
no report about the analysis of phototropism in pinoid mutants.
Interestingly, sequences of PID and PHOT1 kinases show some homology and
they belong to the same AGC kinase family (named after protein kinase A, cyclic
GMP-dependent protein kinases and protein kinase C) (Galván-Ampudia and
Offringa, 2007). Furthermore, PID is interacting with NPY1, which again shows a
homology to NPH3 (Cheng et al., 2007). Cheng et al. hypothesized that the
PID/NPY1 and PHOT1/NPH3 act via similar pathways to modify polar auxin
13
Figure 9. PINOID (PID) affects the development of inflorescence and polar localization of
PIN proteins (Kaplinsky and Barton, 2004).
A: The inflorescence of wild-type Arabidopsis. B: Abscence of PIN1 or PID proteins causes the
same phenotypes, which also appears, when the seedlings are treated by PAT inhibitors. C: Auxin
is produced in young leaves and transported to the root tip, blue arrow depicts the route of PAT.
D: In the shoot, where PID is expressed, PIN proteins have apical localization on the PM. E: In the
root cells, where PID is not expressed normally, PINs are localized on the basal end of the PM,
transport the auxin towards the meristem. F: In shoot of the pid mutant, PIN1 is localized at the
basal pole of cells. This inhibits leaf primordium formation in the shoot. G: In the root ectopically
expressing PID, PIN proteins are localized at the apical poles of cells. This results in decreased
auxin level in meristem, inhibiting the root growth.
transport (Cheng et al., 2007). It is reasonable to postulate that the blue-light induced
relocalization of phototropin1 will affect endosomal processes and auxin transport.
Actually, the PIN2::GFP reporter proteins also change their polar localization
under BL illumination, while red and far red light did not affect it (Laxmi et al., 2008).
In the Chapter 3.3, I will demonstrate the link between BL induced vesicular
movement inside plant cells and polar auxin transport.
Endosomal vesicular movements have been suggested to be involved in the
signal transduction process in animal cells (reviewed by Ibáñez, 2007). Membrane
receptors exist in animal cells, such as the epidermal growth factor receptor (EGFR),
present a receptor-mediated endocytosis (RME), which is a well-known signaling
process in animal cells. It is not a surprising idea to search for the RME process in
14
plant cells. In model plants Arabisopsis, ligand-induced endocytosis of flagellin
receptor was reported (Ali et al., 2007) and also the membrane-localized steroid
receptor kinase BRI1 (brassinosteroid Intensive1) is involved in endosomal signaling
pathways (Geldner et al., 2007). Importantly, BRI1 localizes on the plasma
membrane as homodimers while BRI1 heterodimerizes together with BAK1 (BRI1Associated Receptor Kinase 1) within endosomes (Russinova et al., 2004).
Furthermore, the proteins that are essential for endosomal vesicle transport, such as
SNAREs, Rabs and other proteins, are also found in plant cells. They are playing key
roles in the transduction pathways (Chow et al., 2008, Anders et al., 2008, reviewed
by Samaj et al., 2006). A dynamic motor protein, myosin VIII, has been found to be
involved in the endosomal trafficking in Arabidopsis cells too (Sattarzadeh et al.,
2008). However, the relationships between the cell surface receptor of plant cells and
the intercellular signal transduction are still unclear.
1.6
Introduction to the Drugs Used in this Study
1.6.1 Brefeldin A
Brefeldin A (BFA) is a metabolite of the fungus Eupenicillium brefeldianum. After
its rediscovery in 1988 (Fujiwara et al., 1988), it is now considered a very useful tool
in cell biology, because BFA inhibits exocytosis effectively. This is due to the fact that
BFA prevents vesicle formation in all anterograde exocytosis pathways by stabilizing
the complexes between ADP Ribosylation Factor 1 (ARF1) and the Sec7 domain of
its guanine nucleotide exchange factor (GEF) and such inhibiting the exchange of
phosphate between GTP and the ARF1-GEF-GDP complex. Therefore, the assembly
of coat protein complexes of budding vesicles named COPII is blocked, while the
endocytosis is not affected or even stimulated (reviewed by Chardin et al., 1999). cisGolgi elements merge with the ER in BFA treated cells (Nebenführ et al., 2002),
whereas the trans-Golgi Network (TGN) and endosomes aggregate into so called
BFA-induced compartments. This makes BFA a useful tool for studies of cell biology,
in particular for the study of vesicle trafficking, endocytosis, exocytosis, sorting and
secreting of proteins.
15
1.6.2 Cycloheximide
Cycloheximide (CHX) is an inhibitor of protein biosynthesis in eukaryotes,
produced by the bacterium Streptomyces griseus. It blocks the protein synthesis by
interfering with protein translation, i.e., the translocation of tRNAs and mRNA
molecules on the ribosome (Obrig et al., 1971). By using 35S-methionine as marker of
new synthesis of proteins, I also determined if the conditions of experiments inhibited
protein synthesis completely in root tissues (see the Chapter 2.3.5).
1.6.3 Latrunculin B
Latrunculin B (LB) is a marine toxin, first isolated from the red see sponge,
Latrunculia magnifica. It has been utilized as a powerful drug that rapidly
disassembles actin filaments (F-actin). LB causes F-actin depolymerization by
binding with the actin monomer (G-actin), thereby inhibiting the addition of new Gactin units to the actin filaments (de Oliveira and Mantovani, 1988).
1.6.4 Wortmannin
Wortmannin is a furanosteroid metabolite of fungi Penicillium funiculosum. It is
a specific inhibitor of phosphoinositide 3-kinases (PI3Ks) (Wymann et al., 1996),
involved in the process of endocytosis. Treatment of wortmannin in plant cells inhibits
the vesicular trafficking of membrane proteins from the PM to the endosomal compartment,
the prevacuolar compartment (PVC) and the TGN. Enlarged PVC compartments are
induced by wortmannin (Tse et al., 2004).
1.6.5 FM4-64 (Synaptored) and FM1-43
FM-dyes are nontoxic and water-soluble probes fluorescing only when bound to
membranes. The abbreviation “FM” stands for the chemist’s name, Fei Mao, who
developed the FM-dyes (Ribchester et al., 1994). Both FM1-43 and FM4-64 are
membrane-selective fluorescent dyes. They insert into the outer leaflet of the plasma
membrane lipid bilayer via their lipophilic tails, with the pyridinium dicationic head
anchored at the membrane surface. The amphiphilic nature of these dyes is believed
16
to prevent them from freely crossing from the outer lipid leaflet of the membrane
bilayer to the inner leaflet. The only possible way to internalize the FM dyes is by
endocytosis via the formation of membrane vesicles. FM4-64 and FM1-43 differ
slightly in their chemical structure, and these differences can result in different
patterns of membrane staining and different excitation/emission peaks for the
fluorescence nature (Bolte et al., 2004).
FM4-64 (N-(3-triethylammoniumpropyl)-4-(6-(4-(diethylamino) phenyl) hexatrienyl)
pyridinium dibromide), has excitation/emission peaks at 560/635 nm. The emission
wavelength in the red-domain is a significant advantage compared to FM1-43 (N-(3triethylammoniumpropyl)-4-(4-(diethylamino) styryl) pyridinium dibromide), which
shows a green fluorescence. The red fluorescence of FM4-64 is also advantageous
in studies with GFP-transformed plants (Bolte et al., 2004). SynaptoredTM is the
product name of FM4-64 from company SigmaTM, it is equal to FM4-64.
17
2.
MATERIALS AND METHODS
2.1
Plant Materials
2.1.1 Growth Condition
Seeds of Arabidopsis thaliana were sterilized in 6% NaClO solution with 0.01%
Triton X-100 for 5 minutes. The sterilized seeds were washed with sterilized water for
5 x 1 minute and then planted on agar plates containing 1/2 MS culture medium
(Murashige and Skoog, 1962) and 1% saccharose. Agar plates with seeds were kept
in the refrigerator over night and then placed in the cultivation chamber at 22ºC
temperature and 16 hours illumination per day. Etiolated seedlings were kept in petridishes covered with aluminium foil, after the initiation of germination by 2 hours
illumination with white light at 22ºC.
2.1.2 Genetically Transformed Arabidopsis Lines
1. ProPHOT1::PHOT1::GFP fusion constructs were expressed on the phot1-5
mutant back ground under control of its native promoter (Sakamoto and Briggs,
2002).
2. ProPIN1::PIN1::GFP and proPIN2::PIN2::GFP fusion constructs were also
expressed on the pin1 (pin1-1) and pin2 (eir1-4) mutant background (Delivered from
Rujin Chen, Nobel foundation, USA).
3. ProS35::NPH3::GFP plasmids were made by my colleague Dr. Jan Jasik, the
transformation of this construct into Arabidopsis thaliana is described in Chapter
2.3.1.
4. The proCYCB1::GUS transformed Arabidopsis line (Ferreira et al., 1994), was
used to observe the effects of BL on the root meristem. Cyclin B is a member of the
cyclin family of cell cycle control genes. It binds to the Cdk1 (cyclin-dependent kinase
1). The CYCB/Cdk1 complex promotes expression of genes required during mitosis
(Azzam et al., 2004). Therefore, the activation of cyclin B promoter activity is only
detectable in dividing cells. In plant roots, the presence of dividing cells circumscribes
the region of the root meristem (Hauser and Bauer, 2000). The GUS staining method
will be described in part 2.3.4.
18
5. DR5 is a highly active auxin response promoter element (Ulmasov et al., 1997).
A reporter gene, such as GFP, driven by DR5 is showing the distribution of auxin
signaling, indicating the presence of IAA indirectly. Arabidopsis seedlings were stably
transformed with the proDR5::GFP construct, and by observing the GFPfluorescence, a map of auxin distribution in root tips was established under different
enviromemtal signals.
6. The mutant lines of eir 1-4 (pin2 null mutant, Luschnig et al., 1998) and pin 3-3
(pin3 null mutant, Friml et al., 2002) were used for the phototropic analysis. Both lines
show agravitropic root growth.
7. All Arabidopsis lines were based on the Columbia 0 (Col 0) ecotype except
pin3-3 mutant line, which was based on the Wassilewskija (WS) ecotype.
2.2
Chemicals
In this study, the Arabidopsis seedlings were treated with the inhibitors which are
described in the Chapter 1.6. The concentrations and details of chemical treatments
are shown in the Table 1. The light condition during treatment was carefully
controlled in order to minimize the unwanted light effects from environments. The
controlling of blue light conditions is described in the Chapter 2.3.4.
For labeling of the plasma membrane and endosome compartments, both FM
dyes were used at concentration of 10 µM in 1/2 MS medium. In the experimental
part of Chapter 3.1 and 3.2, FM4-64 and FM1-43 (Molecular Probe ®) are used. In
the Chapter 3.3, synaptored (Sigma ®) was used. Synaptored is the product name of
FM4-64 by Sigma ®. It is chemically equal to the compound originally known as FM464. To avoid quick dye uptake by endocytosis, the seedlings were kept at 4 ˚C for 15
minutes and then treated with the pre-cooled FM working solution at 4˚C for 10
Table 1. List of chemicals.
Name
Storage
Working solution
Company
Brefeldin A
10 mg/ml in DMSO, -20 ºC
50 µM in 1/2 MS
Sigma
Cycloheximide
100 mg/ml in DMSO, 4 ºC
50 µM in 1/2 MS
Sigma
Latrunculin B
2 mM in DMSO, -20 ºC
50 µM in 1/2 MS
Sigma
Wortmannin
10 mg/ml in DMSO, 4 ºC
30 µM in 1/2 MS
Sigma
FM1-43
100 mM in H2O, 4 ºC, dark
10 µM in 1/2 MS
Molecular Probe
FM4-64
100 mM in H2O, 4 ºC, dark
10 µM in 1/2 MS
Molecular Probe
(Synaptored)
(Sigma)
19
minutes. The labeling of shoot epidermal cells needed 60 minutes incubation.
Stepwise labeling of both FM dyes is described in the Chapter 2.3.3.
2.3
Methods
2.3.1 Crossing of transgenic lines of Arabidopsis
1. Transgenetic lines have been planted into soil in the cultivation chamber at 22ºC
temperature and 16 hours illumination per day. After around 4 weeks of growth, when
plants started to develop inflorescences, the largest buds of mother plants were
chosen for crossing. In my studies, the mutant line of phot1-5 was chosen as mother
plant to cross with the PIN2::GFP transformed line, and nph3-1 mutation was chosen
as mother plant to cross with the PHOT1::GFP lines (See the Chapter 2.1.2).
2. The inflorescence was fixed under a binocular by hands gently, only the buds in
right sizes were remained by removing the meristem of the chosen inflorescence. All
the petals and sepals were removed by forceps. Emasculated inflorescences were
marked and the plants were let to grow for 2 overnights. Then the pollens on an
anther of a matured flower of father plants were crossed on matured stigmata. The
seeds matured after about 4 weeks.
3. Seeds were germinated and grown in the above described conditions. The
seedlings were sprayed with herbicide Basta (Bayer CropScience, 1:1000 diluted) to
destroy the uncrossed seedlings. These F1 plants were self-pollinated and grown to
the F2 generation. F2 seeds were cultured on the agar surface as described in the
Chapter 2.1.1, the seedlings with phenotype of phot1-5 and nph3-1 were chosen and
grown on soil to get the F3 generation. The phenotype of lines of F3 have been
analyzed statistically to ensure the homozygous mutant background as described in
the Chapter 2.3.5.
2.3.2 Methods to Produce Transgenic Line of Arabidopsis
1. Preparation of Competent Cells of Escherichia coli and Agrobacterium
tumefaciens.
E. coli bacteria were cultured on the 2% agar plate of LB medium (10g/l Tryptone,
5g/l Yeast extraction, 5g/l NaCl, pH7.0) with selective antibiotics overnight at 37 ºC.
20
A single colony was chosen and cultured in liquid LB medium (around 3 ml) overnight
at 37 ºC on a shaker with speed of 280rpm. 1 ml of this culture medium was added
into 100 ml LB medium, incubated at 28 ºC on the shaker until OD600 reached 0.4.
After chilled on ice for 20 minutes, the medium was centrifuged at 4500 rpm for 10
minutes, in order to spin down the cells. The pellets were resuspended into ice cold
TB buffer (10 mM PIPES, 55mM MnCl2, 15 mM CaCl2, 250mM KCl) and incubate for
30 minutes on ice. After spinning down the cells, these were resuspended into TE
buffer (1/10 volumn) again, divided into tubes as 1.6 ml aliquots with 0.4 ml glycerol,
freezed in N2 and stored in -70 ºC.
The method for preparation of competent cells of A. tumefaciens bacterium is
similar to the E. coli, but the YEP medium (1g/l Peptone, 1g/l yeast extraction, 5g/l
beef extract, 5g/l sucrose, 2mM MgCl2, pH 7.0) was used in this case.
2. Transformation of the Competent Cells by Heat Shock
Stored competent cells were taken from freezer and defrosted on ice. 10 µl
plasmid DNA was added into each tube (40 µl), incubated on ice for 1 hour before a
heat shock in waterbath for 45 seconds. Then, the tubes were put on ice again and
incubated for 2 minutes. 0.5 ml prewarmed LB medium (37ºC) was added to the
tubes and incubated at 37 ºC for 45 minutes on a shaker. As the last step, culture
was spread on antibiotic-containing LB medium plate and cultured over night in 37 ºC
overnight. In case of A.tumefaciens, YEP medium was used to instead of LB medium.
3. Isolation of the plasmid DNA from E. coli by miniprep
Single E. coli clonies were chosen from the agar plates, and cultured in 2 ml liquid
medium at 28ºC on shaker with speed of 180 rpm overnight. The cell suspension was
transferred into 1.5 ml tubes. The cells were spinned-down briefly, the suppernant
medium was discarded, and 100µl solution I (25mM Tris, pH 8.0; 50 mM glucose, 10
mM EDTA, added RNAse freshly) was added into each tubes. The cells were
suspended completely, then 200 µl solution II (0.2M NaOH + 1% SDS, mixed freshly)
was added and mixed gently, and incubated for 3 minutes. 150 µl Solution 3 (3M
potassium acetat, pH5.5) was added to each tube and mixed gently, then centrifuged
for 10 minutes on 13000rpm. The suppernant was transferred into new tubes
21
carefully, without any of the pellets. Double volume of pure ethanol was added into
the suppernant, incubated 10 minutes at 4ºC, the mixture was centrifuged for 20
minutes at 13000rpm. The suppernant was discarded, 70% ethanol was used to
wash the pellet twice, and left to dry out on ice.
4. Transformation of the Competent Cells of A. tumefaciens by Heat Shock
In this step, the plasmid DNA isolated by miniprep was transformed into
Agrobacterium tumefaciens. The methods were same as described in the step 2,
except that the YEP medium was used to instead of the LB medium.
5. Transformation of Arabidopsis plants by Agrobacterium
The plants to transform were planted at the condition described in the Chapter
2.1.3. The first inflorescence was cut in order to remove apical dominance. 4-5
inflorescences were developed in 4-5 days, then the plants were ready for
transformation.
The culture medium (YEP) with transformed Agrobacterium was collected into 50
ml falcon tubes and centrifuged at 3000 rpm for 5 minutes. The cells were
resuspended in infiltration medium (1/2 MS liquid medium, 5% Sucrose, 1mM
Benzylamino Purine (BAP), 0.01% vitamin B5 and 0.01% Silwel) of the same volume.
The inflorescences of the target plants were put into the infiltration media upside
down, incubate in vacuum for 20 minutes for each transformation.
After vacuum infiltration, the inflorescences were inserted into plastic bags and
grown in the culture chamber for two days. Then the bags were removed and the
plants were grown for 4 more days before the vacuum infiltration was repeated once
again to get better transformation rate.
2.3.3 Stepwise Staining of Membrane Fractions in FM Dyes
Seedlings were pre-cooled at 4 ˚C before staining with FM1-43 solutions (5 µM in
the ½ MS medium) at 4˚C for 5 minutes. The stained seedlings were kept in the 1/2
MS medium at room temperature for 5 minutes then incubated with FM4-64 solution (5
µM in the ½ MS medium) at 4˚C for 5 minutes. After a wash with the ½ MS medium
22
for 5 minutes, the seedlings were treated by BFA solution (50 µM) in the dark or
under BL illumination.
2.3.4 Confocal Laser Scanning Microscopy and Blue-Light Illumination
The first two parts of my studies were conducted at the Carnegie Institution
(Department of Plant Biology, San Francisco), with a Biorad MRC1024 confocal laser
scanning microscope, using a Nikon 60x water immersion objective, NA = 1.3.
Excitation of GFP was achieved with the 488 nm laser line of the Ar/Kr-mixed gas
laser. With the Biorad 30% transmittance neutral density filter in place, the fluence
rate at the sample with the 63x water-immersion objective lens was 25 µmol·m-2·s-1.
Fluorescence emission was detected between 505 and 530 nm for PHOT1::GFP
fluorescence and at 650 nm for FM4-64 fluorescence. The external light source used
in some experiments was a halogen lamp (Phillips 20 MR 16, New Jersey, USA)
passed through Corning glass filters: Corning number 5032 for blue light, number
4015 for green light, and number 2404 for red light.
During the third part of my studies microscopy was conducted at the Institut für
physikalische und theoretische Chemie of the University of Bonn, which is equipped
with a LSM 510 meta confocal microscope (Zeiss, Germany), a 60x oil - immersion
objective was used for detailed images and 20x air objective for overviews. An Argon
laser was used to produce the 488 nm blue laser line. Another adjustable BL source
for continous BL illumination came from a halogen bulb. A filter glass (Knight Optical®
with Cat. Number 476FCS5050) was used to provide peak wavelength at 475nm.
The blue laser intensities were measured with a powermeter model 841-PE
(Newport®) equipped with a detector model 818-3T.
2.3.5 Analysis of Whole Seedling Fluorescence
Fluorescence images of whole seedlings were obtained with a Leitz MZFLIII
binocular combined with a digital camera (Leica JVC KY-F708, Leitz,Wetzlau,
Germany). Relative fluorescence intensities were measured with ImageJ software
(Wayne Rasband, National Institutes of Health, Bethesda, Maryland, USA). Images
were background subtracted and intensities of the different regions of the seedlings
were measured for 10 seedlings and averaged for each region. The ‘re-slice’ function
23
of Image J allowed me to construct the cross-section images shown in result Figs. 3
and 5. The dimensions were calculated from the original z-series slices.
2.3.6 Histochemical ß-Glucoronidase (GUS) Staining
Seedlings of stably transformed promoter-GUS plants were stained for ßGlucoronidase activity. Samples were vacuum infiltrated for 10 min with substrate
solution (100 mM sodium phosphate buffer, pH 7.0, 10 mM EDTA, 0.1% Triton X-100,
0.5 mM potassium ferricyanide, 0.5 mM potassium ferrocyanide, and 1 mM 5-bromo4-chloro-3-indolyl glucuronide) and incubated at 37°C for 2h up to 8h. The stained
seedlings were cleared in absolute ethanol and rehydrated by passing through a
graded ethanol series diluted with H2O. The seedlings were kept in H2O, transferred to
microscope slides, and mounted using an anti-fade mounting medium containing pphenylenediamine. Roots were examined using a Leica MZ FL III binocular equipped
with a CCD camera (see 2.3.3).
2.3.7 Determine the New Protein Synthesis by 35S-Methionine
1.
Treatment by 35S-Methionine and CHX
About 0.1 g plant seedlings were collected into 1.5 ml tubes, 0.5 ml TBS buffer
(150 mM NaCl, 10 mM Tris, pH8.0) was added into it. The seedlings were touched
with the buffer but not soaked in it. 0.05 ml CHX (from 5 mM stock solution) was
added into the TBS and treated for 30 minutes, before add 50 micro curies of
35
S-
Methionine to the buffer. Samples were incubated at room temperature for 30 minutes
before protein extraction.
2.
Protein extraction
The buffer was discarded carefully as radioactive waste. Plant tissues were
grinned in extractions buffer (50 mM Tris, 10 mM EDTA, 0.2% Triton X100, 10 mM
mercaptoethanol, pH 7.2, protease inhibitor cocktail (Sigma) 1:2000 and PMSF
100µM were added freshly). After centrifugation briefly (5000rpm, 5 min, 4˚C), the
suppernatant was collected in new tubes as protein solution.
24
3.
SDS-PAGE
The extracted proteins were separated on the polyacrylamide gel by
electrophoresis in this step. Both the stacking gel and the resolving gel were diluted
from the 40% Acrylamid mixture solution (Acrylamid:Bisacrylamid=37.5:1, Gel 40®
from Rotiphorese®). After solidification of the electrophoresis system with resolving
gel (for 20 ml 8% gel: H2O 10.6 ml, Gel 40® 4ml, 1,5M Tris 5 ml pH8.8, 10%SDS
200µl, 10% ammonium persulfat solution 200µl, 20 µl TEMED) and stacking gel (for
5 ml 4% gel: H2O 3.645 ml, Gel 40® 0.625ml, 1.0M Tris 0.63 ml pH6.8, 10%SDS 50µl,
10% ammonium persulfat solution 50µl, 5µl TEMED), the whole system was put into
1Xelectrophoresis buffer (Tris base 3g, Glycine 14.4g, SDS 1g, add H2O to 1l). The
protein samples were mixed up with 1 volume 2Xloading buffer (1M Tris-HCl pH6.8
1.6 ml, 20% SDS 2ml, glycerol 2 ml, beta-mercaptoethanol 1.0ml, bromophenol blue
0.004 g, add H2O to 10ml), then cooked in the boiling water for 2 minutes before
loading gel. Gels were electrophoresed at 120 V in the electrophoresis buffer for 1
hour. The samples were dried in a gel dyer.
4. Recording of radioactive signals
Dried polyacrylamide gel was stacked with an X-ray film (BioMax MR-1 film,
Kodak®) and was exposed for 1 hour in darkness. This experiment suggested that the
1 hour treatment by 50 µM CHX inhibited the protein synthesis completely.
2.3.8 Phototropic Analysis
Seeds of Arabidopsis plant lines were sterilized and planted as described above
(see the Chapter 2.1.1). Because the seedlings of mutant eir1-4 and pin3-3 lost both
gravitropism and phototropism, the seeds were put on the plates in two paralleled
lines. White fluorescent light was filtered by blue foils to create parallel side
illumination with an intensity of 2 µmol·m-2·s-1 (methods to create and measure the
adjustable blue light source are described in section 2.3.2). This blue light intensity
was sufficient to activate the PHOT1 kinase but not the PHOT2.
In the case of proPHOT1::PHOT1::GFP transformed seedlings, they were
cultured under dark condition for 3 days before side illumination with 2 µmol·m-2·s-1
blue light. After 6 and 12 hours illumination, pictures were taken with a digital camera
25
Kodak Z710, only contrast and brightness of pictures were modified as required to
see the images clearly. The separation angles between the position of the root or
shoot tip and the vertical position were measured by the software ImageJ (NIH, USA).
26
3
Results
3.1
Subcellular Localization of Phototropin 1::
::GFP
in Etiolated Seedlings of
::
Arabidopsis thaliana
PHOT1 is expressed in different tissues of 4-days old dark-grown Arabidopsis
seedlings. Because PHOT1 is a photoreceptor which mediates various responses to
BL such as chloroplast movements, opening of stomata, spread of hypocotyls and
phototropism, the description of its distribution could give clues as to understand the
physiological processes behind these responses.
3.1.1 Phototropic Analysis of proPHOT1::PHOT1::GFP Tansformed Seedlings
Before attempting to relate blue-light-induced changes in PHOT1::GFP to any
physiological responses in wild-type plants, it was necessary to determine the relative
physiological sensitivity of the transgenic seedlings to BL compared to wild-type
seedlings. Therefore, both the wild-type (Col 0) seedlings and the phot1-5 seedlings
expressing the PHOT1::GFP gene driven by the PHOT1 promoter were exposed to 2
µmol·m-2·s-1 unilateral BL for different periods of time to obtain a time course for the
development of curvature in each case. The results are shown in Table 2.
Table 2. Complementation of phototropism in etiolated Arabidopsis hypocotyls by
PHOT1::GFP.
Exposure time (hours)
proPHOT1::PHOT1::GFP
Wild-Type
6
19.0 ± 1.5
40.8 ± 3.7
24
49.7 ± 4.5
74.1 ± 4.1
27
3.1.2 Cellular and Subcellular Localizations of PHOT1::GFP in Etiolated
Seedlings
PHOT1::GFP distribution was examined in whole 4-days old dark-grown seedlings
by fluorescence microscopy. At low magnifications, the strongest signals were found
in the hook and the elongation regions of the hypocotyl, and across the abaxial faces
of the cotyledons (Fig. 10A, cotyledon abaxial face view; Fig. 11, cotyledon edge
view). The fluorescence signal declined concomitantly with elongation of the
hypocotyl and expansion of the cotyledons. Whether this decline is simply the
consequence of dilution through cell enlargement or a reduction in gene expression
is not resolved in this study. Surprisingly, a strong signal arose from the shoot–root
transition region (Figs. 10A and B). This strong signal could arise because the cells
are not elongating and diluting the signal, or because PHOT1::GFP expression is
higher in these cells, or both. Fluorescence declined sharply below this region, but
became considerably stronger in the more apical tissues of the root. The average
brightness of GFP signal detected in different tissues of 10 of 4 days old seedlings
Figure 10. PHOT1::GFP expression in dark-grown Arabidopsis seedlings.
(A) PHOT 1::GFP expression in a 4 days old Arabidopsis seedling. Image taken with a binocular
low-power fluorescence microscope (Bar = 1 mm).
(B) Quantification of the intensities of the PHOT1::GFP fluorescence was obtained with ImageJ
software. Arbitrary units, value for highest reading (hook) set to 100. White arrows in Figure 1A
indicate where measurements were taken. Values in Figure 10 B were averaged from 10
seedlings. For details, see Materials and Methods. Error bars indicate standard error of the mean.
28
was measured and the results are shown in Fig. 10B. Table 3 shows the expression
level of PHOT1::GFP in different cell layers of different tissues. Detailed descriptions
of the subcellular expressions and localizations are shown in the Figs. 11 – 18.
Table 3. Expression level of PHOT1::GFP in various tissues of etiolated Arabidopsis
seedlings.
Epidermis
Cortex
Stele
Cotyledon
moderate
moderate
N/A
Apical hook
moderate
strong
moderate
Mature hypocotyl
weak
moderate
moderate
Transition area
weak
strong
weak
Mature root
none
moderate
weak
Root elongation zone
weak
moderate
Moderate
Root cap
none
N/A
N/A
3.1.3 Distribution in the Cotyledons.
At higher magnification, confocal images resolved major differences in the tissue
and cellular distribution of PHOT1::GFP in various tissues and organs. The strong
signals seen from the cotyledons in the Fig.11 arise both from the epidermis on the
abaxial face and the adjacent underlying mesophyll cells. It is almost undetectable
from the adaxial tissues. As shown in the Fig. 12A, the signal from the epidermis at
the margin of the cotyledon is extremely weak (arrows). However, the fluorescence
from the underlying cell layers is far stronger, particularly on certain walls at right
angles to the cotyledon surface (Fig. 12A). These walls are likely the ones most
recently laid down during cell division that precedes cotyledon expansion. They are
much more common near the cotyledon margin where the marginal meristem was
located than toward the center.
The epidermal cells on the abaxial face of the cotyledon are heavily fluorescent
(Figs. 12A and 12B). However, the signal appears enriched at the anticlinal walls and
weaker or undetectable from the inner and outer periclinal walls (Figs. 12 A, B and D).
Using the re-slice option of ImageJ from z-series scanning data, it is possible to
reconstruct transverse images from composite longitudinal images for the Figs. 12C,
12D (see below). Guard cells from 4 days old seedlings as visualized in median
29
transverse section (Fig. 12C) shows signal predominantly on their anticlinal walls.
These anticlinal walls lies at different angles with respect to the vertical walls, this fact
suggests that the intensity of these fluorescence in not an artefact from the optical
axis of scanning. Individual epidermal cells thus resemble imaginative cookie cutters
(Figs. 12A and 12B). There was weak fluorescence at the walls of the mesophyll cells
facing the epidermis (Figs. 12A and 12B). In 4 days old seedlings, the anticlinal walls
of both guard cells show PHOT1::GFP (Fig. 12B). However, in seedlings 3 days old,
incipient guard cells appears as dark circles with weak fluorescence apparent only
around the periphery (Fig. 12E, arrow, cf. Fig. 12B, arrows). Thus, PHOT1::GFP
expression is delayed in guard cells compared with its expression in adjacent
epidermal cells.
Figure 11. Single confocal optical section showing PHOT1::GFP localization in the cotyledon
and apical hook region.
PHOT 1::GFP is preferentially localized at the epidermal anticlinal walls through the hook to the base
of the cotyledon (arrows). The underlying adjacent cortical cells along the hook also show strong
fluorescence at their outer periclinal walls adjacent to the epidermis and at their anticlinal walls, with
little signal at their inner periclinal walls. Thus, each cell forms a ‘C’ shape. Only the abaxial cell
layers of the cotyledon show strong PHOT1::GFP expression (Bar=100 µm).
30
Figure 12. Distribution of PHOT1::GFP in the cotyledons of 4days old seedlings.
(A) and (B): Projection of the abaxial face of a cotyledon near the cotyledon margin of etiolated seedlings.
The image was constructed from a z-series of optical sections of the surface of the cotyledon to a depth of 30
µm. (A): Epidermal cells at the left margin show only very weak PHOT1::GFP fluorescence, localized largely
to the anticlinal walls in a polar fashion (long arrows). The epidermal anticlinal walls are uniformly labeled and
there is almost no detectable signal from the outer or inner periclinal walls. Note the especially strong signal
from cell plate-like structures in the center of the mesophyll cells. (B): The oblique angle of observation
shows the strong signal on the anticlinal walls of the epidermal cells and the lack of signal at inner or outer
walls. The signal is fairly weak in underlying mesophyll cells except for an occasional strongly fluorescent cell
plate-like structure. Note PHOT1::GFP fluorescence from the two pairs of guard cells where they meet
(arrows). (C) and (D): Computer-reconstructed cross-sectional image of guard cells of etiolated seedlings.
Images were produced with ImageJ software by vertically slicing the original z-series dataset. C: Crosssection of epidermis of 4 days old seedling showing guard cells (top center of image). PHOT1::GFP
fluorescence is strongest at the anticlinal walls of the guard cells. Note that the adjacent epidermal cell to the
right lacks fluorescence along its inner periclinal wall. D: Cross-section of epidermis and first layer of cortical
cells. PHOT1::GFP fluorescence is detectable only on the anticlinal walls of the epidermal cells, the anticlinal
walls of the underlying cortical cells, and the common walls between them. E: Projection of abaxial face of
cotyledon of 3 d old light-grown seedling. Image was constructed from a z-series of images of the surface of
the cotyledon to a depth of 30 µm. Note lack of PHOT1::GFP expression at the inner walls in the guard cells
(arrow). (Bars = 20 µm).
31
Another interesting discovery was the localization of PHOT1 on the membrane
surface of chloroplasts. When the seedlings were grown under white light illumination
in order to allow maturation of chloroplasts, the green fluorescence came only from
the surface of chloroplasts facing outwards (Fig.13), therefore only partly overlapping
with the red autofluorescence from chloroplast, whose emission wave length of
autofluorescence is at red region of spectrum (Bolte et al., 2004). From the top view
of projections, images are rebuilt from z-series scanning pictures (Fig. 13A), the two
fluorescence signals are mostly colocalized together, but the green fluorescence
signals is slightly shifted from the red signals (arrows). Observing this region at
higher magnification by step scanning pictures, surfaces of chloroplasts show
colocalization with the PHOT1::GFP signals, but lost the colocalization in images
derived from optical section taken 2 µm deeper into the cell (Fig. 13B, arrow heads).
Figure 13. Localization of PHOT1 on surface of chloroplasts in mesophyll cells.
A: Projections reconstructed from series of images from surface of leaves to 60 µm deep inside.
B: Single images chosen from this z-series image sequence. Upper: 24 µm, middle: 26µm, down:
28 µm from leave surface. (Bar = 20 µm)
3.1.4 Distribution in the Hook
Epidermal cells in the apical hook region show lower expression than the
underlying cortical cells (Figs. 14A and 14B). The epidermal cells on the margins of
the image show that as with the epidermal cells of the cotyledons, there is weaker
signal at the outer periclinal walls (Fig. 14A, arrows). In longitudinal section, it
appears that it is the end walls of cortical cells and the wall adjacent to the epidermal
32
cells that are most heavily labelled (Fig. 14B, thick arrows). This distribution forms a
‘C’ in longitudinal section with the open side facing inward. The heavy signal marks
the contact between the outer periclinal wall of the cortical cell and the inner periclinal
wall of the adjacent epidermal cell arose exclusively from the cortical cells (Fig. 14B,
dashed arrows). Some PHOT1::GFP fluorescence also marks the deeper cells, likely
pith and developing vascular tissue (Figs. 14A and 14B). Because of light scattering,
the resolution is insufficient to provide more structural detail. Note that the end walls
of these inner cells were generally more heavily labelled than the side walls (Fig.
14B)—a polarity of PHOT1 distribution that occurs to varying degrees throughout the
seedling.
Figure 14. Localization of PHOT1::GFP in the apical hook region.
(A) Projection of optical sections from the surface of the hook to a depth of 50 µm. Note smooth
labeling of surfaces of underlying cortical cells, polar distribution of signal in both epidermal and
cortical cells, with strong fluorescence arising from the region of contact between cortical and
epidermal cells. Outermost epidermal walls weakly labeled (thin arrows).
(B) Single image chosen from z-series images. Note ‘C’-like pattern of cortical cells (thick
arrows). Prevascular tissue is marked weakly, and also shows somewhat polar distribution of
PHOT1::GFP fluorescence. Where there is clear physical separation of cortical cells from
epidermal cells, it is obvious that the strong signal at the common faces arises from the cortical
cells and not the epidermal cells (B, dashed arrows) (Bar = 20 µm).
33
3.1.5 Changes from Hook to Root-Shoot Transition Region as Visualized in
Cross-Section
Transverse images were reconstructed from stacks of longitudinal images in Figs.
15 (1–4), illustrating the distribution of PHOT1::GFP in sections through the hook (1),
elongation zone (2), mature zone (3), and the shoot–root transition region (4). The
pattern of strong fluorescence at the anticlinal walls of the epidermal cells, noted
above for the cotyledon epidermis, persists down the hypocotyl axis. In the hook, the
first layer of cortical cells beneath the epidermis also showed the ‘C’-shaped pattern
with the opening facing inwards, just as was the case in longitudinal section (Fig. 151). Further down in the elongation zone, all of the longitudinal walls of the first row of
cortical cells below the epidermis are labelled and the next layer inwards show the
inward-facing ‘C’-shaped distribution of signal (Fig. 15-2). The pattern of labelling in
the shoot at the transition zone is similar to the pattern seen in the elongation zone
(Fig. 15-4). Unlike to the “C” shape of Fig. 14B and Fig. 15-1, this kind of C-shaped
pattern could be artefact, because the superimposition of the laser scanning may
create the strong signals from parallel cell borders. But the differences of cellular
localization of PHOT1::GFP are presented here clearly.
Figure
15.
Computer-reconstructed
cross-sectional
images of hypocotyl cortex and epidermis.
These images were produced with ImageJ software by
vertically re-slicing the original z-series data set.
1: Hook region. Note absence of PHOT1::GFP at inner
periclinal walls of cortical cells (C), outer periclinal walls of
epidermal cells (E), strong signal on outer periclinal and
anticlinal walls of cortical cells (C).
2: Elongation region. Note continued absence of signal at
outer periclinal walls of epidermal cells (E); first underlying
layer of cortical cells (1st C) with stronger signal on anticlinal
than periclinal walls; and labeling of anticlinal walls of next
cortical layer in (2nd C).
3: Mature hypocotyl. The outer periclinal walls of epidermal
cells show a trace of signal and both outer and inner cortical
cells show strongest signal on anticlinal walls.
4: Shoot side of the shoot/root transition region. Outer
periclinal walls of epidermal cells lacks signals either. The
signals from the first layer of cortex are relative fairly and
uniformly distributed (Bar = 50 µm).
34
3.1.6 Distribution of PHOT1 in Elongating and Mature Hypocotyl Tissues
In both the hypocotyl elongation and maturation zones, as in the hook, the
epidermal signal is considerably weaker than that in the cortical cells (Figs. 16A and
B). The pattern of heavy signal seen at the outer periclinal wall and the anticlinal
walls of the cortical cells adjacent to the epidermis of the hook, forming a ‘C’ in
longitudinal section, is not as distinct in the elongation zone, although the ‘C’ shape is
still evident (Fig. 16A, arrows). In the elongating and mature regions, the cortical cells
had moderate signal on all faces (Fig. 16B). A tendency toward bipolar distribution of
the signal remains there. A strong polarity appears in cortical cells, epidermal cells,
and in the stele cells. As in the hook, some GFP fluorescence is present in the inner
tissues (Fig. 16B). These inner cells, possibly vascular parenchyma, show a
tendency toward polar distribution of signal (Fig. 16B, arrows).
Figure 16. Localization of PHOT1::GFP in the elongation zone.
A: Projection of optical sections from the surface of the hook to a depth of 80 µm. Epidermal cells
(weaker signal) show uniform distribution on their anticlinal and outer periclinal walls; cortical cells
(stronger signal) show more uniform labeling, i.e., less polar distribution on all of their walls than in
the hook region. Hence, the ‘C’ pattern of signal distribution is weaker than in the hook (arrows).
B: Single image chosen from the z-series of Fig. 16A. Epidermal cells are uniformly labeled and
the inner periclinal walls of the outer cortical cells are now also labeled so that the ‘C’ pattern is
weaker (arrows). The developing vascular strand (center) shows fairly heavy signal with a
suggestion of stronger fluorescence at the end walls. (Bar=20µm)
35
3.1.7 Distribution at the Shoot–Root Transition Zone
As noted above, there is strong expression of green fluorescence detectable in
the shoot–root transition zone (Fig. 10A). Whether the strong signals reflect a real
increase in PHOT1::GFP expression in this region or simply appear, because these
cells have not elongated as much as their counterparts above them, or both, is an
open question. Both cortical and epidermal cells on the shoot side show the same
pattern of fluorescence as the hypocotyl cells above them—- weaker in the epidermis
than in the cortex, with only slight polarity in both cases (Figs. 17A and B). The
transition from shoot to root is marked by a dramatic and abrupt overall decrease in
signal. Note that GFP fluorescence is completely absent from the root epidermal cells.
The elongated root cortical cells show only very weak signal, largely concentrated at
their end walls in a highly polar fashion (Fig. 17, arrows).
Figure 17. Localization of PHOT1::GFP at the shoot–root transition zone.
A: Projection of optical sections from the surface of the hook to a depth of 100 µm. Cortical (strong
fluorescence) and epidermal (weak fluorescence) cells on the shoot side are relatively uniformly
labeled on all walls. Cortical cells on the root side, by contrast, show sharp polar distribution
(arrows).
B: Single median image from z-series showing somewhat polar PHOT1::GFP distribution in
vascular strand in addition to that in cortical cells. Note complete absence of PHOT1::GFP
expression in the root epidermis (Bar = 100 µm).
36
3.1.8 Distribution of PHOT1::GFP in Mature and Elongating Root Tissues
The expression pattern of PHOT1::GFP in root tissues is different from the pattern
in the shoot region. The epidermal cells of roots of 4 days old etiolated seedlings lack
any detectable expression of PHOT1::GFP. The absence of PHOT1::GFP in the root
epidermis is even more dramatically illustrated in images of roots stained with the
red-fluorescing membrane marker FM4-64 (Fig. 18A–D). Not surprisingly, root hairs,
of epidermal origin, also fail to express PHOT1::GFP (Fig.18B, arrows). The root
cortical cells show polar distribution of PHOT1::GFP through the mature zone and
into the elongation zone (Fig. 18B). The central stele is also labelled as indicated by
the diffuse fluorescence seen in all four images in Fig. 18. Again, technical limitations
(e.g. light scattering and self-shading) prevented resolving individual cell types. The
elongating cortical cells closer to the root tip also show a strong bipolar distribution of
signal (Figs. 18B–D). Along the whole plant bodies, all the cortical and central stele
tissues have expression of PHOT1::GFP, which show strong polar localization in
cells.
3.1.9 Subcellular Distribution of PHOT1::GFP in the Root Apex
As shown in the Fig. 18C, there is relatively strong signal in the elongating region
basal to the root cap. In the more mature region of the root tip, the most intense
fluorescence arises from the stele (Fig. 18B). The red fluorescence from the
membrane stain FM4-64 clearly defines the epidermis in the Figs. 18B–D, verifying a
lack of detectable GFP signal from these cells. As shown in the Fig. 18D, signal is
also undetectable within either the root cap or root apical meristem in these 4 days
old seedlings. Signal first appears in cells destined to become cortex and endodermis,
and only appears more basally in the inner prospective stelar tissues (Figs. 18C and
D). In parallel with the absence of signal at any face of the epidermal cells, there is
only a trace of signal at the outer periclinal walls of the cortical cells. However, their
inner periclinal walls adjacent to incipient endodermal cells share strong signal with
the adjacent endodermal cells and their anticlinal walls are also strongly labelled,
giving them once again the appearance of a ‘C-form’, but this time with the open side
facing outwards (Figs. 18C and D). The smaller endodermal cells also show a ‘C’shaped pattern, sharing heavy signal with the cortical cells and additional signal
37
Figure 18. Distribution of PHOT1::GFP in roots of 4 days old etiolated seedlings.
A: Single low-magnification confocal section of mature root-hair zone. Note heavy expression of
PHOT1::GFP along stele and polar distribution of signal in cortical cells, indicated by colocalization of PHOT1::GFP (green) and FM4-64 (red) resulting in a yellow color. Root hairs
labeled only with FM4-64 are faintly visible (thick arrows). Am, apical meristem; Co, cortex; En,
endodermis; Rc, root cap; St, stele (Bar = 100 µm).
B: Single low-magnification confocal section of root apex. Note heavy PHOT1::GFP expression in
prevascular tissue and cortex, persisting in vascular region and showing distinct polar distribution
in cortical cells. Epidermis shows signal only from FM4-64 and not from PHOT1::GFP (Bar=100
µm).
C: Single high-magnification confocal section of the root apex elongation zone. Note lack of the
PHOT1::GFP signal in epidermal cells (FM4-64 red fluorescence only). Cortical cells heavily
labeled at periclinal walls, only slightly labeled on the side toward the epidermis. Endodermal cells
heavily labeled on the anticlinal walls, slightly labeled on the inner periclinal walls adjacent to stele.
Whether heavy signal between endodermal and cortical cells arises from cortical or endodermal
cells or both could not be resolved (Bar = 20 µm).
D: Single high-magnification confocal section of the extreme root tip. Labeling of cortex and
endodermis precedes labeling of the prospective stelar tissue. Note the complete absence of any
detectable PHOT1::GFP fluorescence from the root cap and apical meristem. The first detectable
fluorescence appears in elongating cortical and endodermal cells (Bar = 20 µm).
38
at their anticlinal walls. There is only weak signal at their inner periclinal walls
adjacent to the prestelar tissue. Hence, the opening of their ‘C’ faces is located
inwards. At this time, it has not been possible to determine, whether the signal along
the common walls between cortex and endodermis is of endodermal or cortical origin,
or both.
39
3.2
BL-Induced Vesicular Re-localization of PHOT1::GFP Molecules
It has already been reported that BL illumination induces relocalization of
PHOT1::GFP from the PM into the cytoplasm (Sakamoto and Briggs, 2002, Knieb et
al., 2004). But the mechanisms and the pathways of this localization are still poorly
understood. Here, the phenomenon is investigated at high resolution in a range of
cell types in cotyledons, hypocotyls, and roots. I have also investigated the minimum
requirement to induce re-localization and the onset of relocalization as a function of
total blue-light fluence rate. Preliminary experiments showed that neither red nor
green light induced any re-localization.
3.2.1 Hypocotyl Cells
The blue-light-induced changes in PHOT1::GFP localization appear as dramatic
reorganization at the cell cortex and the fluorescence signal in the cytoplasm is
clearly detected internal to the cell wall. Figures 19A – C show changes occurring at
the cell membrane of several cortical cells in the hypocotyl elongation zone during
their exposure to the BL from the scanning laser (25 µmol·m-2·s-1). At time zero,
PHOT1::GFP is smoothly distributed over the entire cell surface. The line down the
middle represents the wall of an unlabelled epidermal cell overlying the cortical cell
(Fig. 19A). After 10 min of continuous BL treatment from the confocal microscope
scanning laser, the uniformity once seen at the cell surface is drastically altered and
the signal take on a mottled distribution with distinct dark areas outlined by more
intense signal (Fig. 19B, arrows) and punctuate structures become evident. After 60
min, the mottling increases and more punctuate structures and dark circles are
clearly visible (Fig.19C).
These dramatic changes are accompanied by detection of PHOT1::GFP in the
cytoplasm. Figures 19D–F show a single section through the outermost cell layers
(epidermis and cortex) from the elongating region of the hypocotyl. Again, the bluelight source was the confocal-microscope laser. Already, after 5 min of scanning with
BL, what appear to be strands and blebs of PHOT1::GFP fluorescence are visible in
the cytoplasm in both epidermal and outer and inner cortical cells (Fig. 19E, arrows).
At 10 min, further changes in the intracellular distribution of signal in the two cell
types are visible (Fig.19F).
40
Figure 19. Blue-Light-induced changes in PHOT1::GFP distribution in cortical cells.
Changes in PHOT1::GFP distribution in cortical cells from the elongation region of the hypocotyl
with time of blue-light treatment.
A–C: Image projection showing changes on the plasma membrane at the outer periclinal cell wall of
a cortical cell during blue light treatment. The line down the center cell of A is likely the contact area
to a neighboring cell out of the scanned field of depth. Arrows indicate circular areas devoid of
signal.
D–F: Single images through cortical cells in the same region showing movement of PHOT1::GFP
fluorescence into the cytoplasm. e, epidermis; oc, outer cortex; ic, inner cortex. Arrows indicate
-2
-1
PHOT1::GFP in cytoplasm. Blue-light source: scanning laser (25 µmol·m ·s ). (Bar = 20µm)
Induction of these distributional changes of PHOT1::GFP is very sensitive to BL.
The transgenic seedlings were exposed to three different intensities of BL for 10 min
prior to observation in the confocal microscope immediately thereafter. A total fluence
rate 3 of 300 µmol·m-2 was insufficient to induce any detectable change within that
time period (Fig. 20A). However, 600 µmol·m-2 was sufficient to induce major
changes in PHOT1::GFP distribution (Fig. 20B, arrows) and 3,000 µmol·m-2 likewise
induced dramatic changes (Fig. 20C, arrows). Figure 14A provides a suitable dark
control.
When seedlings were exposed to BL for 30 min (10 µmol·m-2·s-1) and then left in
darkness for 1 h, there was still major reorganization of PHOT1::GFP in cortical cells
in the hook region (Fig. 21A). Empty areas and small punctate structures are clearly
41
Figure 20. Sensitivity of PHOT1::GFP re-localization in hypocotyl cortical cells to blue light
t.
Top view of etiolated seedlings exposed to blue light for 10 min at 0.5 (A), 1 (B), or 5 (C)
-2
µmol·m ·s
-1
-2
(total fluences 300, 600, and 3000 µmol·m , respectively). Projection images were
reconstructed from z-series until 60 µm deep from surface by software. The line down the center
cell of A is likely the contact area to a neighboring cell out of the scanned field of depth. Note
circular areas devoid of signal in B and C (arrows). Blue-light source, halogen lamp plus blue filter
(Bar = 20 µm).
evident in this image (arrows). However, after 2 hours of darkness (Fig. 21B), the
signal assumed a more uniform distribution in several cells (thin arrows), although a
few punctate structures and empty areas (thick arrows) persisted.
The time of onset of the re-localization process is a function of the total photon
fluence of the initial blue-light stimulus (Fig. 22). Dark-grown seedlings were given
total photon fluences ranging from 100 to 10,000 µmol·m-2, all given over a period of
100 s. A new seedling selected from each treatment was examined every 5 min in
the confocal microscope to detect the first evidence for re-localization. When the total
photon fluence was only 100 µmol·m-2, no significant response had occurred after
100 min (Fig. 22A). At 10 times the fluence, re-localization was first detectable after
45 min (Fig. 22B, arrow); at 20 times the fluence, after 20 min (Fig. 22C, arrow); and
after 100 times the fluence, after 5 min (Fig. 22D, arrow). Hence, the greater the
initial fluence was, the earlier the onset of the reorganization. The time of onset
decreased approximately with the log of fluence, suggesting that the response was
the consequence of a first-order photoreaction.
42
Figure 21. Disappearance of PHOT1::GFP from cytoplasm of hypocotyl
cortical cells with time in the dark.
Brightest point projections. Etiolated seedlings were exposed to blue light (10
-2
-1
µmol·m ·s ) for 30 min before being returned to darkness.
A: Appearance after 1 h darkness. Some mottling and dark circular areas
(arrows) still visible.
B: Appearance after 2 h. Signal at many cell surfaces mostly smooth (thin
arrows), although a few empty circular areas persist (thick arrows). Blue-light
source, halogen lamp plus blue filter (Bar=20 µm).
Figure 22. Blue-Light-Induced re-localization of PHOT1::GFP in hypocotyl cortical cells
-2
is sensitive to total photon fluence: (A) 100, (B) 1,000, (C) 2,000, (D) 10,000 µmol m .
Time in dark refers to time in darkness between blue-light pulse and observation in the confocal
microscope. Brightest point projections. Etiolated seedlings were exposed to blue light of
various intensities for 100 s then returned to darkness. At 5-min intervals, hypocotyl cortical
cells were examined in the confocal microscope for evidence of PHOT1::GFP re-localization
(arrows).
43
3.2.2 Cotyledon Cells
Blue-light-activated PHOT1::GFP re-localization is also found in both the
epidermal and mesophyll cells of the cotyledon. The Fig. 23 shows images of the
surface near the cotyledon margin in single optical sections (A–C). As early as 6 min
after the beginning of scanning (25 µmol·m-2·s-1), changes are visible in both cell
types. The surfaces of the underlying mesophyll cells show the beginning of some
mottling and there are small particulate structures within the epidermal cells (Fig.23B).
After 12 min of scanning, there is further release of PHOT1::GFP signal into the
cytoplasm of the epidermal cells (Fig. 23C) and the mottling on the upper surface of
the mesophyll cells had increased.
Figure 23. Blue-light-induced re-localization of PHOT1::GFP in cotyledon epidermal and
mesophyll cells.
(A–C) Projective view of mesophyll cells and epidermal cells. Images were reconstructed from
series of single optical sections scanning through 40µm depth under the epidermal surface. (D-F)
Single optical sections through epidermal cells, chosen from same z-series. Three time points of
illumination by the laser during scanning were chosen as: (A, D) zero time; (B, E) 6 min of
scanning; (C, F) 12 min of scanning. These images show appearance of signal in the cytoplasm of
epidermal cells with time (arrows in E, F), also the stimulation of GFP signals on the mesophyll
PM as punctate structures (arrows in C). Blue light source was the laser light, which illuminated
the specimens during confocal microscopic observation. In the inserts, two pairs of guard cells are
shown to demonstrate that in guard cells PHOT1 did not relocate (Bar = 20 µm).
44
Examination of guard cells on the cotyledons of 4 days old etiolated seedlings
shows that they are unique among the cell types examined in that they did not
undergo a similar re-localization of PHOT1::GFP following blue-light treatment (Fig.
23 D-E). It might be that these guard cells were not fully functional at this stage of
development. Alternatively, light-activated release of PHOT1::GFP into the cytoplasm
simply may not occur in this cell type.
3.2.3 Root Cortical Cells
Because the PHOT1::GFP signal is weaker in the root tissues than in those of the
hypocotyl and cotyledon and PHOT1::GFP is not localized on the outer surface of the
root cortical cells either, it is not possible to see changes that might be occurring at
the plasma membrane itself, comparable to those shown in Figs. 19-22. However,
overall movement of the signal into the cytoplasm following blue-light treatment is
easily observed. Figure 24A–C documents the changes in distribution of green
fluorescence after 0, 6, and 12 min following excitation with the scanning laser. The
signal moves through what appeared to be cytoplasmic strands that converge on a
layer of cytoplasm surrounding the nuclear region (Fig24. arrows in B, C). Under
higher magnification (Fig. 24D), some small punctate structures can be observed
(arrows).
The sensitivity of root cells to BL appears similar to that found in hypocotyl cortical
cells (Fig. 24E–G). Without blue-light treatment, no cytoplasmic PHOT1::GFP is
detectable (Fig. 24E). Ten minutes of exposure to a total photon fluence of 600
µmol·m-2 is sufficient to induce small changes (Fig. 24F). However, 3000 µmol m-2
induced clearly detectable migration of signal into the cytoplasm (Fig. 24G, arrows).
As in the case with the hypocotyl cortical cells, cytoplasmic PHOT1::GFP in root
cortical cells disappears in darkness following blue-light treatment. As shown in Fig.
24 D, the cytoplasm shows little signal after 1 h of darkness following a 30-min bluelight exposure (10 µmol·m-2·s-1). In root cortical cells, dark recovery appears
somewhat more rapid than in hypocotyl cortical cells (Fig. 21).
45
Figure 24. Blue-light-induced PHOT1::GFP re-localization in root cortical cells.
A–C: Zero, 6, and 12 min of blue-light exposure, respectively. Blue-light source is the
-2
-1
scanning laser (25 µmol·m ·s ). (D) High-magnification image showing punctuate structures
in cytoplasm. Note heavily labeled cross walls as compared to longitudinal walls (A, D) and
signal surrounding the nuclear region (B, C). Blue-light source is the scanning laser (25
-2
-1
µmol·m ·s ).
E–G: Sensitivity of root cortical cells to blue light. Etiolated seedlings were exposed to blue
-2
light for 10 min at 0.5 (E), 1 (F), or 5 (G) µmol·m ·s
-1
(total fluences 300, 600, and 3000
-2
µmol·m , respectively). Arrows in G indicate cytoplasmic PHOT1::GFP. Blue-light source is a
halogen lamp fitted with a blue filter (Bar = 20 µm).
3.2.4 BL-Induced Movement and Dark Recovery in the Presence of a Protein
Synthesis Inhibitor
Roots of 4 days old etiolated seedlings were treated with 50 µM cycloheximide
(CHX) for 30 min in darkness prior to observation in the confocal microscope. This
concentration, administered to 3 day old etiolated seedlings for 30 min, completely
prevented the immediately subsequent incorporation of
35
S-methionine into protein
(data not shown). Figure 25A shows that the inhibition of protein synthesis does not
cause any changes of PHOT1 localization in root cortical cells. Migration of
PHOT1::GFP into the cytoplasm in root cortical cells in the elongation region is
clearly evident after 10 min of scanning (BL, 25 µmol·m-2·s-1) (Fig. 25B, arrows) and
even more dramatic after 30 min (Fig. 25C, arrows). Hence, the re-localization itself
46
Figure 25. Effect of cycloheximide (CHX) on blue light-induced PHOT1::GFP re-localization and
subsequent recovery in darkness.
A: Root cortical cells after 30 min CHX treatment in darkness. B: Root cortical cells after 30 min CHX treatment in
-2
-1
darkness and 10 min blue-light treatment at 10 µmol·m ·s . Cytoplasmic PHOT1::GFP is faintly visible. C:
-2
-1
Root cortical cells after 30 min CHX treatment and 30 min blue-light treatment at 10 µmol·m ·s . Cytoplasmic
PHOT1::GFP clearly visible, especially surrounding the nuclear region (arrows, B, C). D–F: Root cortical cells
-2
-1
following 30 min of blue light (10 µmol·m ·s ) and subsequent incubation in darkness in the absence of CHX
(D: 60 min) or in the presence of CHX (E: 60 min, F: 120 min). Traces of signal after 2 h (F) in the presence of
the inhibitor indicate that the disappearance of signal in root cortical cells is considerably slower in the
presence of CHX than in its absence. Light source: halogen lamp plus blue filter (Bar = 20 µm).
brought on by BL does not require protein synthesis. When the roots were kept in
darkness following 30 min of blue-light treatment (10 µmol·m-2·s-1) for 1h (Fig. 25D),
in the absence of CHX, PHOT1::GFP gradually disappeared from the cytoplasm. In
the presence of CHX, the signal also disappeared (Fig. 25E, 1 h darkness; Fig. 25F,
2h darkness). Hence, the disappearance from the cytoplasm might not require
protein synthesis. Whether the disappearance results from return of the cytoplasmic
PHOT1::GFP to the membrane or its degradation can not be resolved at this time,
more experiments are designed to solve this question in next chapters. However, the
disappearance from the cytoplasm (Figs. 25 E and F) is considerably slower than in
the absence of the inhibitor (Fig. 25D).
47
3.2.5
BL-induced Internalization of Membrane-associated PHOT1 via
Endocytosis
Endocytosis of PHOT1 was analyzed with the Zeiss 510-Meta laser scanning
microscope in short time intervals (3.7 seconds per frame with 1.3 sec interval time).
Figure 26 shows that PHOT1 is released from the plasma membrane via a vesiclebased process. The internalized portions of the PHOT1::GFP signals are colocalized
with signals from the plasma membrane (PM) / endosome membrane marker, FM4-64
(Fig. 26). Because of limited passage of FM4-64 through the epidermal cells in to
deeper cell layers, only epidermal cells in the hypocotyl region can be analyzed, but
not cortical cells. In previous study, relocalizations of the PHOT1::GFP in epidermal
and cortical cells showed similar behavior, when the membrane-associated portions
of PHOT1::GFP were traced as mobile structures which were mobilized under
illumination in the cortica cells too (See the Chapter 3.1).
Figure 26. The Released PHOT1::GFP signals from PM of shoot epidermal cells is colocalized with
endosome tracer FM4-64
4-day-old dark-grown seedlings were pre-treated with FM4-64 solution for 60 minutes in dark condition
then put under the confocal laser scanning microscope. Duration of each scan is 3.7 seconds with 1.3
seconds interval time, total time for each scanning is 5 seconds. This sequence starts at the 5th minute of
observation, the yellow arrows heads Figure out a vesicle releasing from cross-wall PM during the
scanning. (Bar = 20 µm)
48
+
Figure 27. The Released PHOT1::GFP signals from the PM of root cortical cells co-localize
with endosomes.
With pre-treatment of synaptored solution for 10 minutes in the dark, the roots of PHOT1::GFP
transformed seedlings were scanned by blue laser for 15 minutes. The series starts at 10 minutes
after beginning of observation, time for each scanning is 3.7 sec with 1.3 seconds interval time.
total time for each frame 5 seconds. Yellow arrows point at an endosomal vesicle released from
cross-wall membrane between two root cortical cells, while blue arrows show synaptored positive
vesicles with less dynamic behavior. (Bar = 20 µm)
In addition, BL-induced punctate PHOT1::GFP-positive structures are scored also
in the root cortex cells (Fig. 27). Again, BL-activated PHOT1::GFP colocalizes with
FM4-64 -positive endosomes in the cytoplasm (arrows in the Fig. 27).
3.2.6 Endosomal PHOT1 Undergoes BL-Sensitive Vesicular Recycling
Brefeldin A (BFA) treatments trapped internalized PHOT1::GFP molecules within
enlarged endosomal compartments in all three cell types analyzed in this study: shoot
epidermis and cortex cells, as well as root cortex cells. Interestingly, these aggregates of
enlarged endosomes, known as BFA-induced compartments, were formed in diverse
cell types at different speed (Table 4). In darkness, the BFA exposure of 10 minutes
was enough for a shift of the PHOT1::GFP-signal into the BFA-compartments while it
49
took about 50 minutes in shoot epidermis cells. In contrast, no compartments were
detected for up to 135 minutes in the shoot cortex cells. BL illumination increased the
speed of the formation of these BFA-compartments in all three cell types (Table 4).
For example, shoot epidermal cells started to show PHOT1::GFP-containing
compartments after 30 minutes of BFA treatment under BL illumination, and even the
shoot cortex cells, which for more than 2 hours did not show any PHOT1::GFP
positive compartments, formed them after 60 minutes (Table 4).
Table 4 . Speed of BFA induced compartment formation in different cell
types (showing the fastest case from each kind of cells, n>3).
Cell Type
Blue-Light/BFA
Dark/BFA
Shoot epidermal
30 minutes
50 minutes
Shoot cortical
60 minutes
No detectable unit 135 minutes
Root cortical
8 minutes
10 minutes
Labelling with FM4-64 reveals endosomal colocalization with the PHOT1::GFP
signals within the same BFA-induced compartments under darkness (Fig. 28). The
regions positive for the PHOT1::GFP signal are larger than those positive for the FM464-signal (Figs. 28, 29), indicating that part of the cytoplasmic PHOT1::GFP signal is
associated with structures or compartments in the immediate vicinity of endosomes.
The time-lapse series (Fig. 29) document the process of BFA-induced trapping of
endosomal membranes coinciding with the internalization of PHOT1::GFP molecules into
the same endosomal compartments. The traces of PHOT1::GFP show more diffused
pattern than the concentrated FM4-64-positive compartments, through this time series,
PHOT1::GFP become more concentrated. Because of the photo-bleach effect from blue
light laser, the whole process of trapping of PHOT1::GFP signals within concentrated
BFA-induced compartments could not be shown (Fig. 29). With results from Fig. 30, the
complete process to build up BFA-induced compartments are clear: Firstly, They are
trapped within small sub-compartments, which are then aggregating into few large BFA
compartments inside each cell. This phenomenon was repeated with different BL
intensities: applying the same time of treatment, a stronger BL illumination caused a
50
Figure 28. Colocalization of
PHOT1::GFP
and
FM4-64
within BFA-induced endosomal
compartments.
FM4-64 pre-incubated seedlings
were treated with BFA for 60
minutes for hypocotyls (A) and 30
minutes for roots (B) in dark
condition. Both shoot epidermal
cells and root cortical cells were
sensitive to BFA. Arrows indicate
the colocalization sites of FM 464 and PHOT1::GFP signals.
(Bar = 20 µm)
Figure 29. Still images from a
time series of cells observed
during BFA treatment.
PHOT1::GFP (middle column) and
FM4-64 (right hand column) are
becoming trapped within the same
compartments during observation
by
confocal
microscopy.
laser
Time
scanning
lapse
series
shows this process at time points
of 1 minute, 6 minutes, 12 minutes
and 20 minutes after scanning.
Arrows
figure
out
the
colocalization sites of FM4-64 and
PHOT1::GFP signals. (Bar=20µm)
51
bigger size of the PHOT1::GFP enriched compartments, while the number of these
compartments per cell decreases (Figs. 30 A-C).
The inhibitor of endocytosis, wortmannin, also affected the localization of
PHOT1::GFP under dark condition (Figs. 30E-F). Punctate structures appeares inside
cytoplasm after short time treatment (Fig. 30E, 15 minutes) and signals inside
cytoplasma were more diffuse than in the long time treated seedlings (Fig. 30F, 60
minutes). But long time treatment did not cause the formation of bigger compartments
than in the short time treated roots. More dramatic impact had the inhibitor of actin
polymerization, latrunculin B, which fully inhibited the assembly of BFA-induced
compartments (Fig. 30G). In contrast, even full inhibition of protein synthesis by pretreatment with cycloheximide (CHX) did not inhibit this process (Fig. 30D).
Figure 30. BFA-sensibility of root cortical cells in different conditions.
A-C show that illumination with higher BL intensity during BFA treatment caused larger phot1::GFP-positive
-
endosomal compartments. The seedlings were treated with BFA (50 µmol l ) under BL illumination with 0 (A),
-2 -
12 (B), 20 (C) µmol m s intensities for 10 minutes.
In order to exclude effects from new-synthesized proteins, seedlings were pretreated for 1 hour by CHX (50)
before incubation in BFA/CHX mixed solution in darknes for 30 minutes, the synthesis of proteins was
stopped totally (see the Chapter 2.3.8). D shows that the formation of BFA-induced endosomal
compartments is partly inhibited, because they are smaller and diffuser than by the BFA treatment without
CHX. The pre-incubation with Latrunculin B can inhibit the formation of BFA-induced compartments totally,
and results in numerous small vesicle-like structures during the same time of BFA treatments (G). The
inhibitor of endocytosis, wortmannin, can also cause the relocalization of PHOT1::GFP molecules, the
pattern of dark-wortmannin treatment for 15 minutes (E) and 60 minutes (F) is shown. (Bar = 20µm)
52
3.3
The Endosomal Recycling of PIN-formed Proteins is Sensitive to the BL
Illumination
PIN proteins are putative facilitators of auxin efflux, and their recycling was
suggested to be an adjustable pathway of polar auxin transport. In the following the
question is addressed as to how the BL signal affects polar auxin transport (PAT) and
how that in turn results in phototropic bending.
3.3.1 PIN2 is Essential for the BL-Induced Phototropic Responses in Roots
The growth direction of the null mutant line of pin2, named as eir 1-4 (Luschnig et
al., 1998) was checked under lateral BL illumination. To this end, the seeds of eir1-4
mutant and WT (Col0) were put onto the same plate and grown under lateral BL
illumination with the intensity of 2 µmol·m-2·s-1. Col0 seedlings had normal phototropic
responses with an average angle for the shoot of 88.9˚ from the vertical position,
while roots had an average angle of 17.5˚. In the eir1 mutant, the lack of PIN2 protein
apparently caused little effects on the shoot phototropic reaction as could be inferred
from the angle of 76.0˚. However, roots of the eir1 mutant lost their gravitropic
responses completely with angles spreading between -38˚~+36˚ (Fig. 31A, n=12).
Furthermore,
roots
of
the
mutant
eir1
seedlings
had
been
Figure 31. PIN2 is essential for root phototropic responses but has no effects in hypocotyl.
-2
-1
Blue light illumination with intensity of 2 µmol·m ·s came from the right side during growth of
80 hrs. The angle of growth directions deviating from the vertical position were measured as γ
(shoot) and θ (root). Because of the zigzag growth of eir 1-4 roots, lines between the root tip and
the initiation points were measured (A, left schema). 12 seedlings were measured in each
Arabidopsis line, ranges of phototropic angels and mean value were recorded (A, right) and
shown schematically (A, middle). B shows several images of both Arabidopsis line, the root parts
are magnified (in white frames).
53
grown in zigzag patterns, irrespective of the light conditions during growth (Fig. 31B).
Even the dark-grown seedlings showed this phenotype (data not shown).
Lack of PIN2 proteins caused nonphototropic responses of Arabidopsis roots. In
order to define wether the polar auxin transport plays essential roles in this process,
the following BFA experiments were designed.
3.3.2 Brefeldin A-inhibited BL-induced Responses in Arabidopsis
In order to study the endosomal cycling in root transition zone cells of Arabidopsis,
and investigate the role of polar auxin transport in phototropic responses, the effect
of Brefeldin A were checked during the light-induced responses in the Arabidopsis
tissues (Fig. 32 and Table 5). The BL signal mediates various responses in plants,
such as phototropism, spreading of leaves, synthesis of chloroplast, inhibition of
hypocotyl elongation and increasing the elongation of root tissues. Figure 32 and
Table 5 suggest that not all of these responses are inhibited by BFA, while the
responses related to auxin are more sensitive than others. The shoot phototropism,
root
elonagtion
and
root
phototropism
have
been
inhibited
or
Figure 32. Brefeldin A inhibits responses to light and gravity.
3-days-old dark-grown Arabidopsis seedlings were chosen for phototropic- and gravitropic
analysis, typical examples are showing as:
A, B: phototropic analysis with blue light from left side for 12 hrs.
C, D: gravitropic analysis in darkness with down side as shown for 12 hrs.
B, C: 500 µM brefeldin A were mixed in the 1/2 MS culture medium
A, D: 1/2 MS medium without BFA.
54
affected, while the Chloroplast development and the expanding of cotyledon remain
in normal status (Table 5).
Table 5. Brefeldin A inhibits responses to light and gravity
(Results are concluded from 15 Arabidopsis seedlings.)
Chloroplast development
Not inhibited
Expanding of cotyledon
Decreased
Stomata opening
Inhibited
Shoot phototropism
Inhibited
Shoot gravitropism
Inhibited
Inhibition of shoot elongation
Root elongation
N/A, (because both BL and BFA can inhibit the
elongation);
Inhibited from 1.3mm/12 hr to 0.3 mm/12 hr
(n=16)
Root phototropism
Inhibited
Root gravitropism
Inhibited
3.3.3 BFA-Sensitive Recycling of PIN2 is Dependent on BL Illumination
Since the endosomal recycling of PIN2 is essential to the polar auxin transport,
the above results suggested that both the lacks of PIN2 protein and inhibition of
protein vesicular secretion could inhibit the phototropism of Arabidopsis. More
experiments were preformed to understand the relationship between the light
illumination and PIN2 recycling. These results are shown in Fig. 33.
Data presented in the Figs. 33A and B suggest that the dark-grown (B) and lightgrown (A) Arabidopsis seedlings have different sensitivities to BFA. After 15 minutes
treatment under white light illumination, BFA treatment caused more and larger
PIN2::GFP positive compartments in cells of the light-grown seedlings (Fig. 33A,
arrows) than those in cells of dark-grown seedlings (Fig. 33B, arrow heads). In darkgrown seedlings, BFA treatments under dark condition did not cause the disappearance
of the PIN2-positive vacuole structures in cells (stars in Fig. 33C-2, comparing to Fig. 35
A), while the FM4-64-positive compartments (arrows in Fig. 33C-1) co-existed without
colocalization with the PIN2::GFP compartments. In contrast, BFA treatments done
under BL illumination resulted in different patterns when both the PIN2::GFP and
55
FM4-64-positive compartments colocalized within same compartments (arrow heads
in Fig. 33D1 and D2).
Figure 33. BFA-sensitive recycling of PIN2 is depended on blue light illumination.
A and B are showing the relocalization of PIN2::GFP in root cells of 4-days seedlings after treated
by 50 µM BFA in light condition for 15 minutes. Seedling in image A was grown under white light
illumination while seedling in image B was grown in dark condition. C and D are showing 4-days
old dark-grown PIN2::GFP transformed seedlings treated by 50 µM BFA in dark condition (C) and
under blue light illumination (D) for 60 minutes, red fluorescence signals are from synaptored. In
dark-BFA treated seedlings, PIN2::GFP (green signals) remains in vacuole structures (defined by
Laxmi et al., 2008) without co-localization with the synapto-red signals. While blue light
illumination causes the dissappearence of PIN2::GFP positive vacuole structures, it involved colocalization of both fluorescence signals inside same BFA-compartments. Arrows are pointing out
the BFA-induced compartments; stars are for the vacuole structures. Bar = 20 µm
56
3.3.4 BL Influences the Localization of PIN 1 and PIN2 Protein.
Since the inhibitor of protein secretion, BFA, stopped gravitropic responses in root
tissues (Fig. 32) and BL affected the sensitivities of root cells to BFA (Fig. 33), it was
reasonable to check, if BL affected localization of PINs via the endosomal vesicle
recycling.
Figure 34 shows the localization of PIN1::GFP proteins in root tips in the dark (Fig.
34A) and the light (Fig. 34B) conditions. PIN1 is expressed mainly in central cylinder
cells, in which they have polar localization in the light-grown plants. Figure 34B
shows that the PIN1::GFP signals are stimulated at the cross-wall membrane, only
weak signal is at side walls and the GFP signals absences in cytoplasm. However, in
dark-grown plants, another pattern of PIN1::GFP localization is prevalent (Fig. 34A).
The PIN1::GFP signal is weaker than in the light-grown plant and it does not show
polar localization. In contract, the signal is found in vacuole-like structures (arrows).
These differences of localizations could be reversed by changes of the growth
conditions. When dark-grown seedlings were placed in white light for 12 hours, the
polarity and the expression levels recovered (Fig. 34C), and no more signal was
found in the vacuole-like structures. If the light grown PIN1::GFP-transformed plants
were put into darkness for 12 hours, the GFP signals reappeared in the vacuole-like
structures (Fig. 34D arrows).
Current hypothesis suggests that, whereas PIN1 acts as auxin transporter in the
central cylinder cells in the direction towards the root apex, PIN2 protein is localized
on the cross wall membrane of epidermal and cortical root cells, in which auxin is
pumped upwards, i.e., away from the apex (see the Chapter 1.2). Therefore the
localization of PIN2::GFP protein was checked in seedlings grown under dark and
light conditions (Fig. 35). Figure 35A shows the localization of PIN2::GFP molecules
in the dark-grown seeding. Both the cross wall membranes and the vacuole-like
structures are labelled in dark-grown seedlings, whereas only the polar membranes
were labelled in the light-grown seedlings (Fig. 35B). These localization patterns can
be reversed too (Figs. 35C and D).
57
Figure
34.
PIN1::GFP
localization and relocalization
in root cells under light and
dark signals.
A: Root of 4 days-old dark-grown
Arabidopsis
seedling,
point
the
at
arrows
vacuole-like
structures. B: Root of 4 days-old
light-grown Arabidopsis seedling.
C: Root of 4 days-old dark-grown
Arabidopsis seedling, which was
transferred to light for the last 12
hours.
D:
localization
of
PIN1::GFP signals in root of 4
days-old light-grown Arabidopsis
seedling, which was transferred
to darknessthe for the last 12
hours.
Arrows
point
at
the
vacuole-like structures (Bar =
20µm).
58
Figure 35. PIN2::GFP localization and relocalization in root cells under light and dark
conditions.
A: Root of 4 days-old dark-grown Arabidopsis seedling, inset shows detail of vacuole-like
structures. B: Root of 4 days-old light-grown Arabidopsis seedling, vacuole-like structures are
absent. C: Relocalization of PIN2::GFP signals in root of 4 days-old dark-grown Arabidopsis
seedling, which had been placed in light for the last 12 hours. D: Relocalization of PIN2::GFP
signals in root of 4 days-old light-grown Arabidopsis seedling, which had been placed in
darkness for the last 12 hours. (Bar = 20 µm)
59
3.3.5 Endocytosis and Fusion of Endosomal Vesicles are Accelerated by BL
Illumination
In order to study the effects of BL on endocytosis and endosomal behavior,
seedlings were stained stepwise with two different sterol dyes for PM and
endosomes: FM1-43 and FM 4-64 (see Materials and Methods). This was combined
with BFA treatments in order to visualize better the changes of endosomal recycling.
Firstly, the question was addressed as to whether FM1-43 and FM 4-64 stained
different portion of the endosomal membrane fractions. When the seedlings were
treated in solution with both dyes together, these two fluorescence signals showed
very similar patterns of localization in root cells of seedlings grown either in dark or in
light conditions (Figs. 36A and B). However, BFA-treatment revealed that the FM 4-64labelled membranes had a more dynamic character than the FM1-43 positive
membranes (Figs. 36 A-1 and A-2). Although BFA treatment caused trapping of both
tracers within the same compartments, more green fluorescence (FM1-43) remained on
the plasma membrane while stronger red fluorescence (FM4-64) was trapped within
the BFA-induced compartments in darkness. Under BL illumination even more
pronounced differences were evident between these two stains: red FM4-64 signal
came mainly from the BFA-compartments, leaving only very weak staining on the
plasma membrane (Fig. 36B-1), while the green FM1-43 signal shows more
prominent plasma membrane labelling (Fig. 36 B-2).
In order to understand better these BL-mediated effects on the endocytosis and
endosome recycling in general, FM1-43 was applied to pre-cooled seedlings
incubated on ice for 5 minutes, then washed with liquid ½ MS medium for 10 minutes
at room temperature. This should cause FM1-43 to be released from the plasma
membrane by endocytosis into the endosomal compartment (early and late
endosomes together). Next, FM4-64 was applied for 5min, which should only reveal
the fraction of early endosomes. All staining steps were performed in darkness. After
BFA treatments for relative long time (60 minutes) in both dark and light conditions, the
FM1-43 and FM 4-64 positive endosomes were all fused together in BFA
compartments (data not shown). However, when the time of BFA treatment was
reduced to 15 minutes, only part of the early endosomes fused together. Using these
incubation parameters under dark condition, fusion of endosomes was relatively low
60
Figure 36. Sequential membrane labelling reveals an effect of light on the fusion of early
and late endosome under BFA treatment.
4 days-old dark-grown seedlings were treated with synaptored (red signal) and FM1-43 (green
signal) mixture (A, B) or treated with synaptored and FM 1-43 sequentially(C, D). Then the
seedlings were treated by 50 µM BFA in darkness (A, C) or under blue light illumination (B, D).
A, B: These images show that both dyes were trapped within same BFA-compartments
(30minutes), when the roots are treated in mixed solution. Synaptored tends to label endosomal
vesicles, while FM1-43 is more strongly on the PMs (Compare A1 A2). Blue light illumination
during the BFA treatment enlarged this difference (Compare B1 B2). Blue light illumination also
increased the BFA-induced endosomal compartments (Compare A and B).
C, D: In order to ensure the different of time-separated endosomal pathways, the dyes were
subjected sequentially and BFA treatments were shortened to 15 minutes. C: 15 minutes BFA
treatment in darkness caused some of the stains to be trapped in the same compartment (yellow
color shows co-localization of two stains), while FM 1-43 (green color) was present in additional
compartments (white arrows); D: 15 minutes BFA treatment in the light resulted in a larger amount
of coloclalization, i.e., only few exclusively green colored compartments could be found.
Bar = 20 µm
(Fig. 36C) whereas, under BL illumination, the same BFA treatment resulted in a much
stronger fusion of endosomes (Fig. 36D).
Furthermore, the obtained pattern of FM 4-64 positive endosomal membrane fraction
revealed also that the BL illumination accelerates general endocytosis. After 30 minutes
of BFA treatment under BL illumination, only low level of the red fluorescence signal
remained at the plasma membrane while most of the signal was trapped within the
61
enlarged BFA-induced compartments (Fig. 36 B-1). On the other hand, BFA treatment
under dark condition resulted in well-labelled plasma membrane and small BFA-induced
compartments (Fig. 36 A-1).
3.3.6 NPH3::GFP has Polar Localization at Root Tip Region
NPH3 is a scaffold protein related to photoreceptor PHOT1 and a homologous to
regulator of PIN localization, PID (see Introduction, Chapter 1.4 and 1.5). In order to
understand relationships between blue light induced relocalization of PHOT1 and
PIN2, I created an Arabidopsis line that was transformed by proS35::NPH3::GFP
constructor (see the Chapter 2.1.2). The preliminary studies are presented as Fig. 37.
In epidermal cells, NPH3::GFP signal was polarly arranged with the crosswalls
showing the strong signals.
Figure 37. Cellular localization of NPH3::GFP in the root tip region.
Figures show the localization of NPH3::GFP in the root epidermal cells of 4 days old light-grown
Arabidopsis seedlings. A: single scanning image from root epidermal cells; B: Projections
reconstructed from series of images from surface of leaves to 30 µm deep inside. Arrows are
pointing the location of NPH3::GFP on the cross wall of two cells. Bar = 20 µm
62
3.3.7 Illumination Changes the Size of Root Meristem and Transition Zone
The size of the apical root meristem was analyzed in proCYCB1::GUS
transformed plant lines under dark and light conditions (see the Chapter 2.1.2).
Under the control of the CYCB1 promotor, the expression of GUS gene pointed out
the region of root meristem in different experiment conditions. When the Arabidopsis
seedlings were grown under light condition, the length of the meristem and transition
zones together was 134.6 µm (mean value from 6 seedlings, Fig. 38 A, B), but in the
dark-grown seedlings it was only 78.2 µm (mean value from 6 seedlings, Fig. 38 C,
D). The dark-grown seedlings only had a basic level of cyclin B promoter-drived GUS
gene expression in this region, whereas seedlings grown under BL illumination had
increased levels of GUS expression. The illuminated side of the roots always showed
a stronger signal than the shaded side (more dark blue colored cells), and the length
of cyclin B expression domain at the illuminated side gets longer than at the shaded
side (Fig. 38E, red arrows). The changes in the light-grown seedlings are not so
significant. But a line of dark blue colored cells was detected at the illuminated
Figure 38. Expression pattern of proCYC B1::GUS in the root tip region.
A and B: Seedlings are grown under white light illumination for 4 days. The length of the
expression region is 134.6 µm (mean value from 6 seedlings).
C and D: Seedlings are grown in dark condition for 4 days. The length of the expression region is
78.2 µm (mean value from 6 seedlings).
E: A dark-grown seedling with 1 hour blue light illumination from the left side (blue arrows).
F: A light-grown seedling with 1 hour blue light illumination from the left side (blue arrows).
Double dead arrows are indicating the length of root meristem, and red arrows are indicating the
strongly expressing cells at the illuminated side of roots. (Bar = 200 µm)
63
side too (Fig. 38 F, red arrows). Figures 38 E and F are chosen from more than 10
seedlings for each experiment, most of them (about 70%) show the same changes.
3.3.8 Asymmetric Auxin Distribution is Affected by Lateral BL illumination
As shown above (see the Chapter 3.3.1-3.3.3) BL illumination determines the
localization and trafficking of PIN2 proteins. In order to understand the importance of
this phenomenon, auxin redistribution was analyzed by observing the GFP
distribution in the proDR5::GFP transformed line of Arabidopsis.
The fluorescence of GFP is detected efficiently at the central column of the root
cap (columella), and the distribution in the columella is symmetrical in the control root
apices grown at diffuse illumination (Fig. 39A). Change of the gravity vector (Fig. 39B)
or 2 hours or unilateral BL illumination with 2 µmol m-2 s-2 from the left side of
seedlings stimulates a shift of the GFP signals towards the illuminated side (Fig. 39C).
Figure 39. Expression pattern of proDR5::GFP at the root tip.
5 days old light-grown seedlings were observed under the confocal microscope. A: Control; B: The
seedling was put into horizontal position for 2 hours. Up and down sides are labelled in this image;
-2 -1
C: The seedling was illuminated with 2 µmol m s blue light for 2 hours from the right side (Arrow
heads). Red arrows point out the asymmetric distribution of GFP signals in root tip. Bar = 50 µm
64
3.3.9 PIN3 is an Essential Factor for Root Phototropism
The importance of PIN3 protein in the root phototropism was demonstrated by
analyzing the phototropic growth of the mutant line pin3-3 (null mutant for the PIN3)
Seeds of pin3-3 mutant and wild type (WS) were put onto the same plate and grown
under lateral BL illumination with the intensity of 2 µmol·m-2·s-1. After 7 days of growth,
images were taken (Fig. 40). It is evident from the zig-zag growth mode, that roots of
the pin3-3 mutant have lost their phototroipsm completely, with growth angles of the
root between -25˚ and +22˚ (12 plants for each line, Fig. 40B). As compared to the
roots of wild type (WS) seedlings, a normal phototropic response with an average
growth angle of 63.0˚ was scored. However, for the shoots of the pin3-3 mutant the
lack of PIN3 caused little effects on the phototropic reactions as can be inferred from
the average growth angle of 42.7˚ as compared to 56.4˚ in the wild type shoots.
Figure 40. Analysis of phototropism in pin3-3 mutants.
Wild type Arabidopsis seedlings have stronger hypocotyl phototropism (56.4º) than the pin3-3
mutant (42.7º). The method of measurement is shown as scheme A, results are in part B. The
mutant loses the root phototropism completely (only 2.2º as mean value), showing a zigzag
formation of root growth (Arrow heads in image C). Schematic diagram C is showing the
distribution of angles of hypocotyl and root growth.
65
4
Discussion
This study was intended to make progress in answering an “old” question: how
can plants, which obviously lack ‘eyes’, sense light signals from the environment?
Especially, how do blue light signals mediate phototropism? Since Charles Darwin
began to study this phenomenon some 150 years ago (Darwin, 1880), researchers
have suggested several hypotheses. Among them, the Went & Cholodny hypothesis
is most widely accepted. Went and Cholodny suggested that the asymmetric
distribution of auxin causes differential cell elongation rates at shaded and
illuminated sides of plant organs (reviewed by Christie, 2007). This then causes the
light-induced bending of the plant organs: hence phototropism. Here, the relocalization
of auxin transporters of the PIN family was observed after blue light perception and
this process is proposed to be one of the key events of phototropism.
Light signals which have their wavelengths in blue range have the most important
roles in the phototropic responses. In 1995, family of blue light receptors were
discovered by Winslow Briggs and his colleagues, and renamed them as PHOT1 and
PHOT2 at 2002. PHOT1 and PHOT2 mediate most BL-intiated responses, such as
phototropism, stomata opening, leaf expansion and chloroplast movement, but the
mechanism of the signal transduction is still unclear (reviewed by Christie, 2007).
Sakamoto and Briggs (2002) published a PHOT1::GFP transformed line of Arabidopsis
plant and suggested new clues to understand the relationship between localization of
PHOT1 and the PHOT1-mediated blue light signaling.
Based on these premier results and hypothesis, I observed in details subcellular
localizations of PHOT1 in different cell types of various tissues with confocal microscopy.
I have discovered that the blue light caused endosomal recycling of PHOT1 between
PM and endosome. In the last part of my thesis, I am suggesting that the blue light
affects the endosomal vesicular trafficking of PINs. This process could determine the
direction of polar auxin transport and drive the phototropic bendings of plant organs.
4.1
The tissue distribution pattern of PHOT1 correlates with Physiological
Responses
In the first part of this study (see the Chapter 3.1), the tissue expression and
subcellular localization of PHOT1::GFP proteins in different kinds of plant cells has
been described. These data support the conclusion that a link exists between
66
localization and function. In the following Chapter it will be discussed how the
distribution of PHOT1 in different tissues is precisely consistent with its physiological
role.
Before attempting to relate blue-light-induced changes in PHOT1::GFP to any
physiological responses in wild-type plants, it was necessary to determine the relative
physiological sensitivity of the transgenic seedlings to BL compared to wild-type
seedlings. The results shown in the Table 2 suggest that the level of expression of
the proPHOT1::PHOT1::GFP gene is either insufficient for full complementation of
phototropism in the phot1-5 mutant or the GFP-fusion protein is not fully functional.
Expression of the GFP-tagged PHOT1 protein is nevertheless sufficient for at least
partial PHOT1 function and to allow confocal examination of light-induced changes in
PHOT1::GFP distribution and re-localization.
4.1.1 PHOT1::GFP Tissue Distribution Patterns with Respect to Physiological
Responses
In the 4-days old dark-grown seedlings, PHOT1::GFP is localized in guard cells of
the cotyledons and in the surrounding epidermal pavement cells. This result is
consistent with the finding that PHOT1 mediates stomata opening (Kinoshita et al.,
2001), and that pavement cells are likely the limiting factor in leaf expansion (Van
Volkenburgh, 1999), respectively. The strong expression of phot1-GFP in the
hypocotyls hook and elongation zone and in the root elongation zone are both to be
expected, because these tissues have prominent roles in the phototropic response
(Schwartz and Koller, 1980, Cho et al., 1996). Moreover, the distribution of phot1GFP in mesophyll cells is consistent with its role in mediating the chloroplast
avoidance response (Jarillo et al., 2001; Kagawa et al., 2001) and accumulation
(Sakai et al., 2001) responses.
Early studies on the shoot phototropism of plants demonstrated the effects of
apical hook position and cotyledon. When the side of hook is positioned away from
the tropic curvature, the bending degree is maximized (Khurana et al., 1989).
However, the authors did not see the possibility that these two organs had roles in
light perception, they only addressed the question as to how their position affected
the phototropic responses. By measuring the Ca2+ flux under blue light illumination
Babourina et al. (2003) came to the conclusion that the light sensor site in the
67
hypocotyl is not limited to the hook region, although the top region of hypocotyl has
the highest sensibility to blue light. This is consistent with the findings here, which
show that the apical hook and elongation zone of hypocotyls have the strongest
expression level of proPHOT1::PHOT1::GFP. PHOT1::GFP is localized on the outer
periclinal PMs. These expression and localization data fit to the reaction site of shoot
phototropism, i.e., the elongation zones (Sakamoto and Briggs, 2002). In summary,
the PHOT1 distribution shown here corroborates all other data on apical hook
opening and cotyledon unfolding stimulated by weak BL (Liscum and Hangarter,
1993).
Sakamoto and Briggs (2002) have noted a lack of detectable expression of
PHOT1 in the root cap as well as the root apical meristem, although they reported a
faint signal from the epidermis. In the present study, this observation was mostly
confirmed except that PHOT1::GFP expression was also not detectable in the root
epidermis. A difference in seedling age or growth conditions might account for this
discrepancy. Historic publications on the identification of the site of root
photosensitivity for phototropism reflect a complex situation. For instance, Naundorf
suggested that the root cap of sun flower seedlings was the site of light signal
perception at 1940, but Schneider (1964) reported that removing of the first 1mm tip
of maize roots inhibited the negative phototropism of root. Mullen et al. (2002)
disagreed with this result, and presented evidences that the root cap of maize
seedlings is the site of photosensitivity for root phototropism. In their studies, white
light applied via optic fibers to the root tip of mays induced curvature, whereas white
light applied similarly to the elongation zone failed to produce a response. They
suggested that the perception and response sites of root are separated. On the other
hand, Liscum and Briggs (1995) demonstrated that it was PHOT1 that mediated
phototropism in response to low-fluence BL in Arabidopsis roots. My analysis reveal
that PHOT1::GFP is absent from the root cap and apical meristem in 4 days old
etiolated seedlings. Either Arabidopsis and maize use different systems to regulate
root phototropism or another photoreceptor is responsible for photoexcitation in the
root cap. A gradient of light intensity between the illuminated and the shaded sides of
a plant would be the physical basis for plant bodies to sense the direction of light.
Optical fibers were used to measure the light gradient of the plant vessel tissues
irradiated by unilateral BL. Light penetration, known to occur in etiolated tissues
(Mandoli and Briggs, 1982) and light scattering (Vogelmann and Haupt, 1985), could
68
result in a light gradient in the elongation zone, even if only the root cap were directly
illuminated but could not account for the greatest photosensitivity occurring in the cap
region. This assumption is receiving quite some credibility, as is shown here,
because roots illuminated with 2 µmol·m-2·s-1 of BL clearly curved away from the light
in the absence of detectable PHOT1::GFP fluorescence in their apical meristems or
root caps before or after phototropic stimulation (Fig. 18). As Mullen et al. (2002)
suggest, perhaps the activation of phytochrome, known to be located almost
exclusively in the root-cap columella cells in roots of maize and other etiolated grass
seedlings (Pratt and Coleman, 1974), is required to potentiate a tropic response to
BL in the phototropin-rich elongation region. BL can transform phytochrome in vivo,
although the Pfr:Pr (far red : red) ratio is small and the quantum efficiency low
compared to these parameters in RL (Pratt and Briggs, 1966). Because Mullen et al.
(2002) used continuous BL, phytochrome activation certainly must have occurred.
It is not clear why PHOT1::GFP expression is high in the root–shoot transition
region. Galen et al. (2007) have reported that PHOT1 serves a role in drought
tolerance in field-grown Arabidopsis. Even if the zone is under the soil, penetrating
light could provide sufficient phototropin activation to induce some sorts of response
(Mandoli and Briggs, 1982). Under drought stress, blue-light-activated PHOT1 in the
transition region might send signals to the roots that could modulate their growth rate
or growth direction. Aspects of root phototropism, meristem size and positioning and
PAT under BL illumination will be discussed in further detail in the Chapter 4.3.3.
4.1.2 Subcellular Distribution of PHOT1
With higher-resolution confocal microscopy a great deal better subcellular details
of PHOT1::GFP localization can be observed than that found in the previous study by
Sakamoto and Briggs (2002).
It has been well established for a number of plant species that PHOT1 is closely
associated with the plasma membrane in dark-grown seedlings (Briggs et al., 2001).
Results from the present study are consistent with that view. In dark-grown seedlings,
PHOT1::GFP fluorescence is found closely associated with the outermost surface of
the cytoplasm in all cell types in which we found it expressed. What is surprising is
that different cell types and similar cell types at different stages of development may
show unique patterns. In the cotyledons, distribution appears uniform at the plasma
69
membrane of mesophyll cells but is limited almost entirely to the anticlinal walls of the
epidermal cells. Little, if any, signal was detected on the periclinal walls (Figs. 11, 14,
15). Expression in the marginal epidermal cells was especially weak compared to the
cells on the abaxial surface of the cotyledons (Fig. 12). Like other epidermal cells, the
guard cells also have PHOT1::GFP largely on their anticlinal walls (Fig. 12). However,
it appears later in guard cells than in the other epidermal cells (Fig. 12B and 12E)
and shows particularly strong expression on the contacting walls between the two
guard cells of a single stoma (Fig. 12B). In the dividing tissue at the cotyledon
margins, there is also exceptionally strong expression at what appears to be recently
laid down cell walls of the mesophyll cells (Fig. 12A). Blue light mediates the
expanding of cotyledons and leaves, Van Volkenburgh (1999) suggested that it is the
layer of mesophyll cells increase cotyledon expanding but the epidermal cells limit
the expanding. It is reasonable to suppose that the PHOT1 increase the dividing rare
of mesophyll cells under illumination.More experiments are needed to support this
hypothesis.
PHOT1 also mediates the chloroplast avoidance (Jarillo et al., 2001; Kagawa et
al., 2001) and accumulation (Sakai et al., 2001) responses. The localization of
PHOT1 gives clues to understand these processes. The chloroplasts develop in the
mesophyll cells of 4 days old light-grown seedlings. In these cells, PHOT1::GFP
signals are found on the surface of chloroplasts rather than the PM (Fig. 13). PHOT1
may interact with the J-domain protein, JAC1, in the process of initiating chloroplast
movement (Suetsugu et al.,2005) and it has been shown that motor proteins of the
myosin class VIII are involved in the PHOT1-dependent blue light-stimulated
movement of chloroplast (Krzeszowiec et al., 2007). This suggests that the actin
cytoskeleton plays an important role the PHOT1-mediated intercellular movements.
Actually, disrupting actin filaments can stop the recycling of PHOT1::GFP too (Fig. 30
G).
In the hypocotyle hook and elongation zone, there is strong expression of
PHOT1::GFP localized along the anticlinal and outer periclinal walls of cortical cells,
moderate abundance along the end walls of elongating epidermal cells, and weak
abundance at their outer walls (Figs.14 and 16). These PHOT1::GFP signals are
forming a ‘C’-shaped pattern in longitudinal section, with the strongest signals from
outer periclinal walls. This subcellular localization of PHOT1 reflexes the role of the
hypocotyle hook and elongation zone in the phototropic responses (see the Chapter
70
4.1.1). The localization of PHOT1on the outer membrane provides an efficient pattern
of photoreception. In elongating and mature tissues, as the cell surfaces become
more completely labelled, outer periclinal walls do not have such priority. The polar
localization of PHOT1 on the cross walls suggested that the PHOT1 may play
intracellular roles here.
4.1.3 Cross-Wall Localization of PHOT1 in the Root Transition Zone Suggests
Roles of PHOT1 in Polar Auxin Transport
Unlike the localization of PHOT1 in the shoot, most of the PHOT1::GFP signal is
found along the cross walls in cortical cells of the root elongation region, especially
the root transition zone, whereas the periclinal walls are only weakly labelled in these
cells. These results are comparable to the subcellular localization of PIN2 protein,
which is mainly localized on the cross-wall membranes of cortical and epidermal cells
of the transition zone (Fig. 18 and Fig. 35).
At the root tip, members of PIN-formed proteins build up a transportation network
to modify the distribution of auxin (Blilou et al., 2003). Under gravitropic and
phototropic stimulation, they determine the asymmetry of distribution between both
root sides, resulting in differential elongation of cells on both sides. PIN2 acts as the
regulator for the auxin transport from the root tip upwards. PHOT and PINOID (PID)
proteins have similar evolutionary ancestors and belong to the same protein family,
namely the AGC kinases (Galván-Ampudia and Offringa, 2007). PID is well-known to
affect the function of PIN proteins as pid mutants are losing the apical polar
localization of PIN1 protein (Friml et al., 2004). PID and PIN1 and PIN2 proteins are
partly co-localized at the cross-wall plasma membrane in Arabidopsis roots and
hypocotyls. This observation suggests that PHOT1 and PIN2 proteins may not only
be co-localized on the same plasmamembrane domain but may also functionally
interact. Furthermore, PID is interacting with NPY1, which again shows a homology
to NPH3 (Cheng et al., 2007). Cheng et al. hypothesized that the PID/NPY1 and
PHOT1/NPH3 act via similar pathways to modify polar auxin transport. The result in
Fig.37 supports this hypothesis well, because the localization of NPH3::GFP also
shows perfect polarity, and NPH3::GFP localized all on the cross wall membrane of
root cells. However, this hypothesis still needs more additional experiments to be
verified.
71
The polar localization of PHOT1 in root cells is also one reason, why work in this
thesis is focused on the root tip of Arabidopsis plants, which is not a popular object
for phototropic analysis.
72
4.2
BL-Induced Relocalization of PHOT1::GFP is achieved by Receptor
Mediated Endocytosis
Sakamoto and Briggs (2002) have reported blue-light-induced movement of
PHOT1::GFP into the soluble fraction of the cytoplasm and Knieb et al. (2004) have
obtained a similar result using native PHOT1 protein in solution. In both studies highspeed centrifugation was used to separate membrane and soluble proteins. Knieb et
al. (2004) estimated that at least 20% of the total PHOT1 was released to the
cytoplasm
following
blue-light
treatment
of
etiolated
mustard
seedlings.
Immunoblotting did not provided any evidences for breakdown products in either
study. The question as to whether some of the cytoplasmic PHOT1::GFP is present
within intracellular vesicles has remained unresolved so far, although results reported
by Kong et al. (2006) indicate that PHOT2 appears to migrate into vesicles that colocalize with a Golgi marker upon blue light induction. In the following (Chapter 4.2.2),
data will be presented, which support the localization of PHOT1 within endosomal
vesicles.
4.2.1 Blue-Light Effects on PHOT1::GFP Subcellular Distribution
The reasons for blue-light-induced re-localization of PHOT1 and its movement
into the cytoplasm are not clear. It could be a mechanism to desensitize plant tissues
to BL by removing photoreceptor from the region of the plasma membrane, where it
is required to interact with other PM-associated proteins. As mentioned in the
introduction, BL induces some major ionic changes in plant tissues. It has been
shown by Fuchs et al. (2003), that the potassium channel, ZMK1, is expressed as a
gradient across unilaterally illuminated maize coleoptiles. However, the gradient
developed relatively slowly, only after 60 min irradiation and therefore it is unlikely to
be related directly to blue-light-induced changes in subcellular distribution. Cho and
Spalding (1996) reported a rapid transient depolarization in Arabidopsis hypocotyls
exposed to BL, attributable to the activation of anion channels. This transient
response occurred well before we observed the first changes in PHOT1::GFP
distribution (Figs. 19, 20). Whether these changes bear a causal relation to
subsequent changes in PHOT1 distribution remains to be determined.
73
There are ionic changes with kinetics similar to those of the blue-light-mediated
PHOT1::GFP re-localization and recovery described here. Baum et al. (1999)
reported rapid but transient increases in calcium uptake in response to a blue-light
pulse by light-grown Arabidopsis seedlings expressing the calcium sensing aequorin
system. The overall changes lasted less than 100 s. However, a single blue-light
pulse desensitized the system to subsequent pulses and recovery took place over 3–
4 h, not unlike the dark recovery to a strict localization of PHOT1::GFP at the plasma
membrane reported in the present study (Fig. 22). Two groups have reported bluelight-induced increases in calcium uptake by etiolated tissues of Arabidopsis (Stoelzle
et al., 2003) and several other dicot species (Babourina et al., 2003), respectively.
Stoelzle et al. (2003) found the response strongly reduced in a phot1-5 mutant and
completely absent in a phot1-5/phot2-1 double mutant. In both studies, the kinetic
was consistent with those for blue-light-induced PHOT1::GFP re-localization, with the
changes detectable several minutes after the onset of BL and continuing for 15 or
more minutes thereafter. It would be premature to do more than point out the
correlation of these ionic changes with changes in the subcellular distribution of
PHOT1::GFP. Further experimentation should be performed to determine, whether
there is a causal relationship between these changes in calcium movement and the
loss of PHOT1 from the plasma membrane in BL and its subsequent dark recovery to
a plasma-membrane-only distribution.
4.2.2 Internalization of PHOT1 is Accomplished via Endocytosis
Data presented in section 3.2 showed that internalization of BL-activated PHOT1
sensor occurred in two separate steps. Firstly, PHOT1::GFP accumulates in punctate
structures, which are then released from the plasma membrane and co-localize with
endosomal membranes (Wan et al., 2008). These PHOT1::GFP enriched endocytic
vesicles can be traced by the quick scanning mode of the microscope (Fig. 26). The
PHOT1::GFP signal is firstly assembled into aggregates associated with the plasma
membrane. Subsequently, the PHOT1::GFP positive structures are released as discrete
spots. The membrane endocytic marker FM4-64, which emits red fluorescence in the
hydrophobic environment of the membrane, is a very useful tool for the studies of
endocytosis. FM4-64 can not cross the double lipid layer of the plasma membrane,
and the only way that this membrane tracer can get access to the cytoplasm is via
74
endocytosis. The data in the Figs. 26 and 27 prove that the PHOT1::GFP-positive
vesicles are part of the membrane recycling endocytic system. FM4-64 labelled
endosomes show PHOT1::GFP fluorescence, suggesting that at least part of
PHOT1::GFP internalization is associated with endocytosis.
Prevacuolar compartments (PVC) and trans-Golgi networks (TGN) have a function in
protein secretion and endosomal recycling (reviwed by Šamaj et al., 2005, Jürgens
2004.). In order to characterize the movement of endocytic PHOT1 in plant cells, the
inhibitor of exocytosis and vesicle recycling, brefeldin A (BFA), and the endocytic
inhibitor, wortmannin were applied. The results show, that consistent with other
reports on BFA-action (Nebenfuhr et al., 2002), PHOT1::GFP molecules are trapped
within the enlarged BFA-compartments. This finding provides strong evidence that
internalization of PHOT1 occurrs along endosomal recycling pathways (Fig. 28).
Protein synthesis is not required for this process, since cycloheximide pre-treatment
does not stop the formation of PHOT1::GFP enriched BFA-induced compartments in
dark treated cells (Fig. 30D).
Wortmannin, an inhibitor of PI3 kinase and PI3 kinase related enzymes, targets
the early endosomal transportation in animal and plants cells (Simonsen et al., 1998;
Mills et al., 1999). In plant cells, Wortmannin traps PVC-localized proteins within ringlike structures, while the TGN-localized proteins remain unaffected (Miao et al., 2006).
In addition, Tse et al. (2006) suggested that Wortmannin causes stacking of PVCs
into enlarged endosomal structures, which are enriched also with endocytic PIN2
(Jaillais et al., 2006). Secretion of proteins is not effected by Wortmannin. It is
conceivable that wortmannin affects PHOT1 because of its membrane-bound nature
(Figs. 30E and F). Taken together, the reactions to both inhibitors support the
assumption that PHOT1 is recycled via PM-endosomal-TGN-PVC-PM recycling
pathways. Additional experiments using the actin inhibitor latrunculin B (Fig. 30G)
have proven that this pathway is dependent on the actin cytoskeleton.
4.2.3 ARF1-GTPase, the Key Regulator of Endocytosis, is Modified by PHOT1
ADP-ribosylation factors (ARFs) regulate vesicular trafficking and protein
secretion/sorting in many kinds of organisms at the interfaces between Er and Golgi,
endosomes and PM and possibly between other endomembrane compartments
(Donaldson and Honda, 2005, Anders et al., 2008). GNOM is a plant specific
75
member of the ARF Guanosine Exchange Factor family (ARF-GEF) which activate
ARFs in the processes of protein sorting und vesicle budding. GNOM has also been
shown to modify the sorting and vesicular trafficking of PINs (Anders et al., 2008). As
a key regulator of endocytosis, the functional role of ARF-GEFs is still far from clear.
In Arabidopsis, GNOM regulates the vesicular trafficking between Endosomes and
the Plasma membrane (Teh and Moore, 2007). Once activated by ARF-GEFs, ARFs
recruit coat proteins such as COPI (Sun et al., 2007, Antonny et al., 2005) and
regulate phospholipid metabolism; they also modulate the structure of actin filaments
on the membrane surfaces (Myers and Casanova, 2008).
BFA specifically targets ARF-GEFs (Peyroche et al., 1999). The BFA-sensitivity of
PHOT1 suggests, that the endosomal recycling of PHOT1 is related to the ARFmediated pathways. Uhrig and coworkers (Botanisches Institut of University of
Cologne, Joachim Uhrig, personal communication, unpublished data) analyzed the
protein interaction between plant ARFs (ARF1, ARF6, and ARL1) and PHOT1, 2
proteins. By using the yeast two-hybrid assay, they found a direct linkage between
the active form of ARF 1 (Q71L and д17) and both PHOT1 and PHOT2 proteins. The
inactive form of ARF1 did not show such linkage to PHOT1 (Joachim Uhrig,
unpublished).
Clearly, further studies are needed in order to determine the role of PHOTs in the
interaction with ARFs. It is still unclear, whether the blue light-activated PHOT1
regulates the function of ARF molecules then modify the endosomal recycling, or the
ARF-GEF complex is participating in the recycling of PHOT proteins. The first
hypothesis may have more potential than the second one, because the level of
PHOT1 recycling reflects the intensity of blue light illumination, this process seems to
act as the trigger in the signaling pathways.
4.2.4 Endocytosis of PHOT1::GFP is Affected by BL Illumination
Under BL illumination, part of the PHOT1 molecules is released from the PM via
endocytosis. In Chapters 3.1, it was suggested, that internalization is a two step
process: at first, PHOT1::GFP signal are collected on the cell surface together, then it
moves inside the cytoplasm. The speed and the level of this process reflect the
amount of photons arrived on the surface of the plant. The fate of the internalized
portion of PHOT1 is still unclear. Sakamoto & Briggs (2002) suggested that a part of the
76
internalized PHOT1 is degraded, because the total amount of PHOT1 decreased
under continuous BL illumination. The data collected in this thesis support this
hypothesis, because co-localization of PHOT1::GFP with FM4-64 was not identical
and there were diffuse clouds of PHOT1 around the co-localized endosomal
compartments. Immunoblotting studies also suggested that part of PHOT1 becomes
soluble after illumination (Knieb et al., 2004), and Sakamoto suggested that after 24
hours of illumination, the total amount of PHOT1 proteins is decreased dramatically. All
these studies suggest that the blue-light activated PHOT1::GFP is internalized via two
different endocytic pathways: one targets on the activated PHOT1 for degradation and
the other feeds into the endocytotic vesicular recycling pathway. Alternatively, there
may be only one endocytic pathway and the internalized PHOT1 population would
then be sorted subsequently either for degradation or for endocytic recycling.
Furthermore, the size of BFA-induced endosomal compartments corresponds to
the rate of exocytosis of target proteins. After the inhibition of protein synthesis by
CHX, PHOT1::GFP is still trapped within these BFA-induced compartments in both
dark and light conditions. This fact suggests that the recycling of PHOT1 between the
plasma membrane and endosomal membrane system is active even under dark
conditions. However, the BL illumination increased the rate of PHOT1 recycling, as
inferred from the increased size of the BFA-induced compartments. This is true for all
cell types investigated in the present study (Table 4). Furthermore, increasing the BL
intensity during BFA treatment also stimulates the size of PHOT1::GFP enriched
BFA-induced endosomal compartments (Fig. 30 A-C). Comparing the time lapse
images in Fig. 29, it can be concluded, that BL accelerates the recycling of PHOT1
between endosomes and the plasma membrane. It is very attractive to speculate,
that this endosomal recycling of activated PHOT1 represent an integral part of BLinduced signal transduction.
4.2.5 Signaling Endosomes as Integrators of Environmental Signals?
From our knowledg of cell surface receptors in animal cells, such as receptors for
epidermal growth factor and transferring (reviewed by Ibáñez, 2007), for example, we
can infer that activated receptors will undergo endocytosis in plants too. Receptor
mediated endocytosis (RME) has indeed been documented in plant cells. This was
achieved by observing the uptake of recombinant human transferrin (hTfr) expressed
77
in Arabidopsis (Ortiz-Zapater et al., 2006). Another recent example is the
brassinosteroid receptor, BRI1, which is internalized by the RME process (Geldner et
al., 2007). Furthermore, the proteins that are essential for endosomal vesicle
transport, such as SNAREs, Rabs and other proteins, are also found in plant cells.
They are playing key roles in the transduction pathways (Chow et al., 2008, Anders
et al., 2008, reviewed by Samaj et al., 2006). A dynamic motor protein, myosin VIII,
has been found to be involved in the endosomal trafficking in Arabidopsis cells too. It
proved an actin filament-dependent mechanism of RME in plant cells (Sattarzadeh et
al., 2008). This thesis provides new evidences to understand relationships between
the cell surface receptor of plant cells and the intercellular signal transduction. The
rate of both, internalization as well as endosomal recycling of the BL activated
PHOT1 is positively correlated with strength and duration of illumination. So it is
reasonable to hypothesize, that BL-induced endocytic internalization of the activated
PHOT1 is involved in BL signal perception and/or transduction. In this process, BL
has a ligand-like role and activates the PHOT1 by conformation changes.
4.2.6 Model to Describe the PHOT1-Mediated Signaling by Endosomal
Vesicular Relocalization.
Fig. 41 shows steps of PHOT1 relocalization schematically. Firstly, PHOT1 is
localized on the PM uniformly and smoothly. The changes of molecular conformation
or the status of phosphorylation cause internalisation of PHOT1 into endosomal
vesicles. The mechanism of this process is still unclear. It may be speculated, that
NPH3 acts as a scaffold (Pedmale and Liscum, 2007) and PHOT1 may be
internalized as dimers (Salomon et al., 2004). As the first known component between
endocytosis and BL receptor, ARF1 helps PHOT1 containing endosomal vesicles to
fuse with the TGN or PVC compartments. This process even operates under dark
condition, while blue light signals accelerate the internalisation of PHOT1. The
experiments also suggested that the level of PHOT1 internalization reflexes the
intensity and time of blue light illumination. Furthermore, the exchange of ARF-GEFGDP (inactive form) for ARF-GEF-GTP (active form) is blocked by BFA. Thus, it can
be expected that the recycling of PHOT1 is involved in the endomembrane pathways,
and it accelerates recycling of endosomes (see Fig. 41). ARF-GEF is needed in this
78
process. Does this process have roles in signal transduction? How can it affect the
auxin polar transport and involve in lateral differences in cell elongation?
Figure 41: Model to explain PHOT1 movement under blue light illumination.
1.
Activation of PHOT1 protein by blue light illumination, the activated PHOT1 molecules
are collected at the PM domain of coated pit.
2.
The forming of coated vesicles and internalization into endosomes is affected by
wortmannin. This step is induced by blue light. The activated form of ARF factors supply
docking sites for PHOT1.
3.
Early and late endosomes are involved in the recycling pathway.
4.
Recycling pathways of PHOT1 are sensitive to BFA, which blocks the exocytosis step by
binding on the GNOM then stopping the activation of ARF,
5.
Part of the PHOT1 molecules are recycling between endosomes and PVC
compartments, this process is wortmannin-sensitive. These data suggest part of the
internalized PHOT1 is shuttled between PVC and endosome.
6.
Part (appr. 20%) of PHOT1 becomes soluble in the cytoplasm.
7.
Soluble and vacuolar portion of PHOT1 is degraded. The degradation may be mediated
by the ubiquitin proteasome pathway.
79
4.3 Intracellular Localization and Endosomal Recycling of PIN-formed
Proteins are Determined by BL Illumiation
PIN – formed proteins (PINs) have been considered as auxin efflux carriers or
facilitator ever since they have been discovered. As detailed in the Introduction, the
recent in vivo analysis of auxin flow has drawn out a network of PIN-formed proteins
in the root tip. This determines direction and rate of polar auxin transportation in
response to gravi- and phototropic stimuli. In this network, PIN1 helps polar auxin
transport from shoot to root tip through the cell files in the central cylinder, while the
outer cell layers (cortex and epidermis) transport auxin backwards with the help of
PIN2.
4.3.1 Light Signal Changes Subcellular Localization of PIN1 and PIN2 Proteins
The polar localization of PIN proteins on the cross-wall membrane is well-known,
but only few studies have reported the relationship between polar localization and
light signaling (Blakeslee et al., 2004, Laxmi et al., 2008). Laxmi also included that in
dark condition, the PIN2 was localized in vacuolar compartments efficiently. In this
thesis, data have been presented which allow to conclude that the polar localization
of both PIN1 and PIN2 exist only in the seedlings grown under light condition. In dark
condition, vacuolar compartments were labelled with PIN1::GFP or PIN2::GFP
signals. Blue light illumination caused the disappearance of these vacuole-like
accumulation of PINs (Figs. 34 and 35), Laxmi (2008) discovered that the
disappearance of these compartments was inhibited by inhibitor of ubiquitin
proteasome, this indicated that the vacuole-portion of PINs were destined for
degradation under blue light illumination. In order to exclude the possibility that the
vacuolar portion of PIN2 was removed via exocytosis, the seedlings were treated by
BFA under blue light illumination. In this experiment, FM4-64 was also used to trace
the endomembrane trafficking. When the roots were treated in BFA solution during
the BL illumination, BFA can not inhibit the disappearance of these vacuolar
compartments (Fig. 31). No more PIN2::GFP signals were found inside the vacuolar
compartments. These results supported that the vacuole portion of PINs is going to
the ubiquitim-depended degradation pathways, but not to the BFA-sensitive
80
exocytosis. The physiological function of this vacuolar portion of PIN2 is still unclear.
Even more experiments were needed to discover its ultimate fates.
4.3.2 The BFA-Sensitive Endosomal Recycling of PIN2 is Depeneded on Blue
Light Illumination.
Though the BFA did not inhibit the blue light -depended degradation of PIN2, the
BFA-sensitive vesicular recycling of PIN2 was strongly effected by blue light
illumination. The BFA-treatment under blue light illumination trapped PIN2::GFP
signals within the enlarged endosomal compartments, in which the FM4-64 signals
were colocalized. On other hand, the BFA treatment in dark condition did not cause
the same pattern of PIN2 relocalization and FM4-64 colocalization. In the etiolated
seedlings, formation of vacuolar compartments with PIN2::GFP was not changed by
the dark-BFA treatment. This fact again indicted that the BFA-sensitive exocytosis of
PIN2 was separated from the light-stimulated degradation of PIN2. In order to
exclude all the exocytosis pathways from this PIN2 relocalization, a wide range of
inhibitors need to be applied in the experiments.
BFA is an inhibitor specifically targets ARF-GEFs. BFA specifically targets ARFGEFs. As a result of this action, the recycling of PIN1, PIN2 proteins and polar auxin
transport are inhibited. The polar distribution of PIN proteins is also depended on the
activation of ARF (Kleine-Vehn et al., 2008). Sauer and colleagues (2006) suggested
that the auxin signaling did not effects the transcription of PINs, but the polar
localization of PINs was controlled by auxin molecules via feed back pathways.
As we discussed in Chapter 4.2.3, the ARF factor is also a key factor in blue lightmediated endocytic recycling of PHOT1. The results in my studies reveal logical
linkages between BL signaling and rates of endocytic trafficking directly. Firstly, BL
signals activate the PHOT1 by changing the protein conformation. Amount of PHOT1
internalized into cytoplasm depends on illumination time and on light intensities.
Although the mechanisms are not clear, these dynamic parameters and the
colocalization with FM4-64 suggest that the endocytosis of PHOT1 is accomplished
via a receptor mediated endocytosis. Secondly, the GNOM localization in plant cells
is specific on early/recycling endosomes which are targets of BFA. Therefore,
PHOT1 is trapped within BFA-induced endosomal compartments together with FM464. At the same time, PIN1 and PIN2 proteins are also trapped in the FM4-64
81
Figure 42. Model to explain how blue light modifies PIN-mediated auxin transport.
1. Blue light increases the endocytosis rate of PHOT1.
2. Under dark condition, small vacuoles (i.e., prevacuolar compartments) contain PIN2 which is
released under blue light illumination via unknown pathways. PIN2 is then localized on the
endosomal compartments and supports the efflux of auxin.
3. At least part of the vacuole localized PIN2 was degradated via ubiquitin pathways.
4. PIN2 supports transmembrane transport of auxin molecules. This process does not only exist
across the PM, but also across the membrane of endosomes. Polar auxin transport depended
on polar localization of this vesicular recycling pathway.
5. Vesicle-based export of auxin is BFA sensitive. BFA blocks exocytosis and stops the polar
auxin transport.
enriched BFA-induced compartments. These data suggest that the blue lightactivated PHOT1 can accelerate the recycling rate of PIN proteins and modify the
auxin polar transport in a direct way. Thirdly, the active forms of ARF (ARF-GTP)
have physical relationship with the PHOT1 on endosome compartments. The
activated ARFs are likely to give PHOT1 docking sites at endosomal membranes. BL
82
stimulation increases the trafficking of PHOT1 in the cytoplasm via endocytosis, this
process may affect the vesicle fusion in the trafficking pathways. Furthermore, the
fusion of early and late endosome in BFA treatments are suporting this hypothsis (Fig.
35).
4.3.3 Root Phototropism
In this thesis, experiments were undertaken to elucidate the intracellular
localization and fate of BL photoreceptors in response to illumination. As discussed in
Chapter 4.3.2, that the endosomal trafficking of PHOT1 can modify the PIN2
recycling under BL illumination. Here I analyzed root phototropism of null mutant pin2
(eir1-4). Mutant seedlings lost the abilities of phototropic response to BL signals
completely, while the shoot organ was less influenced due to the lack of PIN2
(Chapter 3.3.1).
In Arabidopsis roots, PIN2 has similar distribution as PHOT1. Interestingly, both
of them are not expressed in the root cap. This is surprising, at least in Zea mays, the
root cap is supposed to be the organ of light perception (Mullen et al., 2002). The
root cap has been suspected as the light perception organ of Arabidopsis too.
However, there is still no direct analysis of this question. The analysis with pin3-3
gives indirect evidences that the PIN3 has essential roles in the phototropic
responses of root apices (Chapter 3.3.9). The relocalization of PIN3 proteins in the
columella root cap cells are observed under reorientation of root in gravity vector, and
has been supposed to be the mechanism to distribute auxin asymmetricly (Friml et al.,
2002). Whether a similar mechanism plays a role under lateral illumination is still
unclear. Moreover, asymmetric auxin distribution at root cap has been observed in
the proDR5::GFP transformed Arabidopsis seedlings too (Chapter 3.3.8). Thus, root
cap is clearly one of the sensory organs for photoreception. Because PHOT1 is not
expressed in root caps, there must be other photoreceptors involved. PHYA and
PHYB can be candidates with high potentials. PHYA and PHYB are essential to the
RL induced root positive phototropism of Arabidopsis plants (Kiss et al., 2003) and
BL can also activate these putative RL receptors and inhibit the gravitropic responses
of Arabidopsis (Lariguet and Frankhauser, 2004). Studies with phyA and phyB
mutants are important to discover the mechanism of root apex phototropisms.
Another clue comes from the studies to analysis the region of meristem. Dark-grown
83
seedlings have smaller meristem than the light gown seedlings. Furthermore, the
illuminated side shows extended and more active cell divisions than the shaded side.
Based on these results, it is clear that the PIN proteins are not only essential for
the root gravitropic responses, but also for the root phototropisms. Asymmetric
distribution of auxin occurs at illuminated root tips of Arabidopsis too. PIN3 is
expressed at root caps and redistributes auxin to the shaded side under lateral light
illumination. The signals of gravity and lateral light are sensed by root cap cells.
Subsequently, the root tip bends to the light source at an angle of about 18 degree
(Fig.40), until the competition of two kinds of signals reaches balance. More studies
are needed to find out which photoreceptor(s) mediate(s) this process.
At the meristem and root apex transition zone, PHOT1 modifies the endosomal
recycling of PIN2 in cortical cells. The recycling of PIN2 adjusts the rate of upward
auxin transport and gives another clue to integrate both signals of gravity and light.
This analysis results in a “delayed” response to light signals. Mullen (2000) found that
the Arabidopsis roots response to light occur at the central elongation zone, while the
roots respond to gravity at the root apex transition zone. Here I suppose that the root
apex transition zone does not only act as reaction organ to gravitropism, but also as
sensory region for integrating different environmental signals. Clearly, more detailed
analysis is needed to improve our still limited understanding how sensory signals
from gravi- sensing and light-sensing are integrated in order to allow all adaptive root
growth behavior.
84
Figure 41 Schematic model to describe how the blue light modifies PAT pathways.
Arrows are showing the PAT pathways in the root tip of Arabidopsis. Left: root in vertical position
without light signals; Right: root in vertical position with lateral blue light illumination (Blue arrows).
Apical transportation of auxin via central cyclinder cells is modified by the auxin efflux facilitators
PIN1 (green arrow). At the root cap, the PIN3 distributes auxin to the shaded side predominantly.
The mechanism of this process is still unclear (Red arrow). PIN2 helps to shift the PAT towards the
epidermis and cortex and PIN3 tansports auxin in basal direction through the epidermis and cortex
(Purple arrow). This transportation pathway is enhanced by the blue light activated receptor
PHOT1 via vesicular trafficking. For the sake of simplicity, I did not include PIN4 and PIN7 which
are also expressed in cells of root tips.
85
5.
Outlook
Based on results of this thesis, new problems are identified and new questions
emerge. For example, it is unclear how PHOT1 and ARF molecules interact via their
interaction domains. The recently published structure of PHOT1 does not give an
answer to this question. Though it is widely accepted that polar localization and
vesicular recycling of PINs is regulated by the ARF-GEFs, the particular molecular
mechanisms are still unclear. Without understanding these mechanisms, it is not easy
to explore further the exact signal transduction pathways linking PHOT1 endocytosis
with the recycling of PINs.
To this end, it is important to conduct a domain analysis of ARF, ARF-GEF and
PHOT1, and introduce point mutations to PHOT1. Of particular interest is the
question wether the phosphorylation site of PHOT1 interacts with ARF. Furthermore,
I have also crossed the PIN2::GFP construct into the phot1-5, phot1-2/phot2-1 and
nph3-1 mutant backgrounds. This will allow identification of those members of the
blue light signaling pathways which modify the PIN2 relocalization.
86
6.
Summary
In this thesis, the subcellular localization and possible functions of the blue light
receptor, PHOT1, is examined. Results are presented, which indicate that blue lightinduced relocalization of auxin efflux carriers (PIN-proteins) is adjusted by PHOT1 via
ARF-mediated and BFA-sensitive vesicle trafficking pathways.
PHOT1 was visualized by expressing the PHOT1::GFP reporter in Arabidopsis
mutants which contain a dysfunctional copy of the endogenous PHOT1-gene. It is
shown that the subcellular localization of PHOT1 in plant cells reflects the
physiological functions and development stages of special kinds of cells. For example
in guard cells, PHOT1 appears only in the mature developmental stage, when these
cells begin to react to blue light signals. Especially, the specific roles of PHOT1 were
addressed in root and shoot apical meristems. The obtained data suggest that
PHOT1 has different roles in root and shoot phototropic responses. PHOT1 in root
tissues acts to adjust the polar auxin transport, while PHOT1 in shoot tissues has
more abundant roles. It is also suggested that the mechanism of phototropic
responses in shoots and roots might be different from each other.
Interestingly, the PHOT1::GFP signals are quickly released from the plasma
membrane and appear in intracellular vesicles, when cells are irradiated with blue
laser light in the process of scanning in the confocal microscope. The level of PHOT1
relocalization reflects the amount of photons reaching the surface of the Arabidopsis
cells. PHOT1-positive vesicles colocalize with the endosomal tracer, FM4-64,
indicating that that light-induced movements of PHOT1-positive endosome are
analogous to the ligand-induced receptor mediated endocytosis. PHOT1-mediated
blue light sensing is sensitive to inhibitors of exocytosis and endocytosis and is also
dependent on intact actin filaments.
Endosomal recycling of PIN proteins is considered as the mechanism to adjust
the levels of polar auxin transport. Here it is demonstrated that blue light affects the
recycling of PIN1 and PIN2 between the plasma membrane and the endosomal
compartment. The pin2 null mutant lacks phototropic responses in roots, while the
shoot phototropism in this mutant is independent on PIN2. By taking into account the
87
polar transport of auxin under asymmetric illumination at the root tip, a testable model
for the mechanism of root phototropism is proposed, which will helpful in guiding
future phototropism research .
88
References
Abas L., Benjamins R., Malenica N., Paciorek T., Wiśniewska J., Moulinier-Anzola
J.C., Sieberer T., Friml J., Luschnig C. (2006) Intracellular trafficking and
proteolysis of the Arabidopsis auxin-efflux facilitator PIN2 are involved in root
gravitropism. Nat Cell Biol. 8, 249-256.
Ali G.S., Prasad K.V., Day I., Reddy A.S. (2007) Ligand-dependent reduction in the
membrane mobility of FLAGELLIN SENSITIVE2, an Arabidopsis receptor-like
kinase. Plant Cell & Physiol. 48, 1601-1611.
Anders N., Nielsen M., Keicher J., Stierhof Y.D., Furutani M., Tasaka M., Skriver K.,
Jürgens G. (2008) Membrane association of the Arabidopsis ARF exchange
factor GNOM involves interaction of conserved domains. Plant Cell. 20, 142-151.
Azzam R., Chen S.L., Shou W., Mah A.S., Alexandru G., Nasmyth K., Annan R.S.,
Carr S.A., Deshaies R.J. (2004) Phosphorylation by cyclin B-Cdk underlies
release of mitotic exit activator Cdc14 from the nucleolus. Science. 305, 516-519.
Babourina O.K.,Newman I.A., Shabala S.N. (2003). Electrophysiological localization
of blue light sensory sites in etiolated dicotyledon seedlings. Plant Cell Environ.
26, 1505–1514.
Baluška F., Šamaj J., Menzel D. (2003) Polar transport of auxin: carrier-mediated flux
across the plasma membrane or neurotransmitter-like secretion? Trends in Cell
Biology. 13, 282-285.
Baluška F., Volkmann D., Menzel D. (2005) Plant synapses: actin-based adhesion
domains for cell-to-cell communication. Trends in Plant Science. 10, 106-111.
Baluška F., Schlicht M., Volkmann D., Mancuso S. (2007) Vesicular secretion of
auxin: Evidences and implications. Plant Signaling & Behavior. 3, 254-256.
Baum G., Long J.C., Jenkins G.I., Trewavas A.J. (1999) Stimulation of the blue light
phototropic receptor NPH1 causes a transient increase in cytosolic Ca2+. Proceedings
of National Academy of Sciences USA. 96, 13554-13559.
Bennett C.L. (2005) Astrophysical observations: lensing and eclipsing Einstein’s
theories. Science. 307, 879-884.
Blakeslee J.J., Bandyopadhyay A., Peer W.A., Makam S.N., Murphy A.S. (2004)
Relocalizatin of the PIN1 auxin efflux facilitator plays a role in phototropic responses.
Plant Physiol. 134, 28-31.
89
Blilou I., Xu J., Wildwater M., Willemsen V., Popper I., Friml J., Heldstra R., Aida M.,
Palme K., Scheres B. (2005) The PIN auxin efflux facilitator network controls growth
and patterning in Arabidopsis roots. Nature. 433, 39-44.
Briggs W.R., Beck C.F., Cashmore A.R., Christie J.M., Hughes J., Jarillo J.A.,
Kagawa T., Kanegae H., Liscum E., Nagatani A., Okada K., Salomon M., Rüdiger
W., Sakai T., Takano M., Wada M., Watson J.C. (2001) The phototropin family of
photoreceptors. Plant Cell. 13, 993-997.
Bonifacino J.S., Jackson C.L. (2003) Endosome-specific localization and function of
the ARF activator GNOM. Cell. 112, 141-2.
Bolte S., Talbot C., Boutte Y., Catrice O., Read N.D., Satiat-Jeunemaitre B. (2004) FMdyes as experimental probes for dissecting vesicle traffickingin living plant cells.
Journal of Microscopy. 214, 159-173.
Briggs W.R., Christie J.M. (2002) Phototropins 1 and 2: versatile plant blue-light
receptors. Trends in Plant Science. 7, 204-210.
Chardin P., McCormick F. (1999)Brefeldin A: the advantage of being uncompetitive.
Cell. 97, 153-155.
Chen R., Masson P.H. (2006) Auxin transport and recycling of PIN proteins in plants.
Endocytosis in Plants. Plant Cell Monography. 1, 139-157.
Cheng Y., Qin G., Dai X., Zhao Y. (2007) NPY1, a BTB-NPH3-like protein, plays a
critical role in auxin-regulated organogenesis in Arabidopsis. Proceedings of
National Academy of Sciences USA. 104, 18825-18829.
Cho M.H., Spalding E.P. (1996) An anion channel in Arabidopsis hypocotyls
activated by blue light. Proceedings of National Academy of Sciences USA. 93,
8134-8138.
Cho H.Y., Tseng T.S., Kaiserli E., Sullivan S., Christie J.M., Briggs W.R. (2007)
Physiological roles of the light, oxygen, or voltage domains of phototropin 1 and 2
in Arabidopsis. Plant Physiology. 143, 517-529.
Cholodny N. (1926) Beiträge zur Analyse der geotropischen Reaktion. Jahrbuch für
Wissenschaftliche Botanik 65: 447–459.
Chow C.M., Neto H., Foucart C., Moore I. (2008) Rab-A2 and Rab-A3 GTPases
define a trans-golgi endosomal membrane domain in Arabidopsis that contributes
substantially to the cell plate. Plant Cell. 20:101-123.
Christie J.M. (2007) Phototropin blue-light receptors. Annual Reviews of Plant
Biology. 58, 21-45.
90
Christensen S.K., Dagenais N., Chory J., Weigel D. (2000) Regulation of auxin
response by the protein kinase PINOID. Cell. 100, 469-478.
Correll M.J., Kiss J.Z. (2002) Interactions between gravitropism and phototropism in
plants. J Plant Growth Regul. 21, 89-101
DARWIN C. (1880) The Power of Movements in Plants (assisted by F. DARWIN).
John Murray, London
De Marais D.J., (2000) Evolution. When did photosynthesis emerge on Earth?
Science. 289, 1703-1705
De Oliveira C.A., Mantovani B. (1988) Latrunculin A is a potent inhibitor of
phagocytosis by macrophages. Life Sci. 43, 1825-1830.
Dhonukshe P., Aniento F., Hwang I., Robinson D.G., Mravec J., Stierhof Y.D., Friml J.
(2007) Clathrin-mediated constitutive endocytosis of PIN auxin efflux carriers in
Arabidopsis. Current Biology. 17, 520-527.
Dyall S.D., Brown M.T., Johnson P.J. (2004) Ancient invasions: from endosymbionts
to organelles. Science. 304, 253-257.
Eitoku T., Nakasone Y., Zikihara K., Matsuoka D., Tokutomi S., Terazima M. (2007)
Photochemical intermediates of Arabidopsis phototropin 2 LOV domains
associated with conformational changes. Journal of Molecular Biology. 371, 12901303.
Esmon C.A., Pedmale U.V., Liscum E. (2005) Plant tropisms: providing the power of
movement to a sessile organism. International Journal of Developmental Biology.
49, 665-674.
Folta K.M., Maruhnich S.A. (2007) Green light: a signal to slow down or stop. J Exp
Bot. 58, 3099-3111.
Folta KM., Spalding EP. (2001) Unexpected roles for cryptochrome 2 and phototropin
revealed by high-resolution analysis of blue light-mediated hypocotyl growth
inhibition. Plant J. 26, 471-478.
Folta K.M., Lieg E.J., Durham T., Spalding E.P. (2003) Primary inhibition of hypocotyl
growth and phototropism depend differently on phototropin-mediated increases in
the cytoplasmic calcium induced by blue light. Plant Physiol. 133, 1464-1470.
Freddolino P.L., Dittrich M., Schulten K. (2006) Dynamic switching mechanisms in
LOV1 and LOV2 domains of plant phototropins. Biophysics Journal. 91, 36303639.
91
Friml J., Wisiewska J., Benkova E., Mendgen K., Palme K. (2002) Lateral relocation
of auxin efflux regulator PIN3 mediates tropism in Arabidopsis. Nature. 415, 806809.
Friml J., Yang X., Michniewicz M., Weijers D., Quint A., Tietz O., Benjamins R., Ouwerkerk
P.B., Ljung K., Sandberg G., Hooykaas P.J., Palme K., Offringa R. (2004) A PINOIDdependent binary switch in apical-basal PIN polar targeting directs auxin efflux.
Science. 306, 862-865.
Fuchs I., Phillipar K., Ljung K., Sandberg G., Hedrich R. (2003) Blue light regulates an
auxin-induced K+-channel gene in the maize coleoptile. Proc Natl Acad Sci U S A.
100, 11795-11800.
Fujiwara T., Oda K., Yokota S., Takatsuki A., Ikehara Y. (1988) Brefeldin A causes
disassembly of the Golgi complex and accumulation of secretory proteins in the
endoplasmic reticulum. J Biol Chem. 263, 18545-18552.
Ferreira P., Hemerly A., de Almeida Engler J., Bergounioux C., Burssens S., Van
Montagu M., Engler G., Inzé D. (1994) Three discrete classes of Arabidopsis
cyclins are expressed during different intervals of the cell cycle. Proc Natl Acad
Sci U S A. 91,11313-11317.
Galván-Ampudia C.S., Offringa R. (2007) Plant evolution: AGC kinases tell the auxin
tale. Trends in Plant Science. 12, 541-547.
Galen, C., Rabenold, J.J., Liscum, E. (2007). Functional ecology of a blue light
photoreceptor: effects of phototropin-1 on root growth enhance drought tolerance
in Arabidopsis thaliana. New Phytol. 173, 91–99.
Geldner N., Friml J., Stierhof Y.D., Jurgens G., Palme K. (2001) Auxin transport
inhibitors block PIN1 cycling and vesicle trafficking. Nature. 413, 425-428
Geldner N., Anders N., Wolters H., Keicher J., Kornberger W., Müller P., Delbarre A.,
Ueda T., Nakano A., Jürgens G. (2003) The Arabidopsis GNOM ARF-GEF
mediates endosomal recycling, auxin transport, and auxin-dependent plant growth.
Cell. 112, 219-230.
Geldner N., Hyman D.L., Wang X., Schumacher K., Chory J. (2007) Endosomal
signaling of plant steroid receptor kinase BRI1. Genes & Development 21,15981602.
Harada A., Sakai T., Okada K. (2003) PHOT and PHOT2 mediate blue light-induced
transient increases in cytosolic Ca2+ differently in Arabidopsis leaves. Proc Natl Acad
Sci U S A. 100, 8583-8588.
92
Hause G., Šamaj J., Menzel D., Baluška F. (2006) Fine structural analysis of brefeldin Ainduced compartment formation after high-pressure freeze fixation of maize root
epidermis: Compound exocytosis resembling cell plate formation during
cytokinesis. Plant Signaling & Behavior 1, 134-139.
Hauser MT., Bauer E. (2000) Histochemical analysis of root meristem activity in
Arabidopsis thaliana using a cyclin:GUS (β-glucuronidase) marker line. Plant and
Soil. 226, 1-10.
Hubert B., Funke G.L. (1937) The phototropism of terrestrial roots. Biol Jaarboek. 4,
286–315.
Ibáñez C.F. (2007) Message in a bottle: long-range retrograde signaling in the
nervous system. Trends Cell Biol. 17, 519-528.
Iino M. (2006) Toward understanding the ecological functions of tropisms: interactions
among and effects of light on tropisms. Current Opinion in Plant Biology. 9, 89-93.
Inoue H., Randazzo PA. (2007) ARF GAPs and their interacting proteins. Traffic. 8,
1465-1475.
Inada S., Ohgishi M., Mayama T., Okada K., Sakai T. (2004) RPT2 is a signal
transducer involved in phototropin response and stomatal opening by association
with phototropin 1 in Arabidopsis thaliana. Plant Cell. 16, 887-896.
Jackson C.L., Casanova J.E. (2000) Turning on ARF: the Sec7 family of guaninenucleotide-exchange factors. Trends in Cell Biology. 10, 60-67.
Jacob T.C., Moss S.J., Jurd, R. (2008) GABAA receptor trafficking and its role in the
dynamic modulation of neuronal inhibition. Nature Reviews Neuroscience. 9, 331343.
Jaillais Y., Fobis-Loisy I., Miège C., Rollin C., Gaude T. (2006) AtSNX1 defines an
endosome for auxin-carrier trafficking in Arabidopsis. Nature. 443, 106-109.
Jarillo J.A., Gabrys H., Capel J., Alonso J.M., Ecker J.R., Cashmore A.R. (2001)
Phototropin-related NPL1 controls chloroplast relocation induced by blue light.
Nature. 410, 952-954.
Kagawa T., Sakai T., Suetsugu N., Oikawa K., Ishiguro S., Kato T., Tabata S., Okada
K., Wada M. (2001) Arabidopsis NPL1: A photoropin homolog controlling the
chloroplast highlight avoidance response. Science. 291, 2138-2141.
Kaplinsky N.J., Barton M.K. (2004) Plant biology. Plant acupuncture: sticking PINs in
the right places. Science. 306, 822-823.
93
Khurana J.P., Best T.R., Poff K.L. (1989) Influence of hook position on phototropic
and gravitropic curvature by etiolated hypocotyls of Arabidopsis thaliana. Plant
Physiol. 90, 376-379.
Kinoshita T., Doi M., Suetsugu N., Kagawa T., Wada M., Shimazaki K. (2001)
PHOT1 and PHOT2 mediate blue light regulation of stomatal opening. Nature 414,
656-660.
Kinoshita T., Emi T., Tominaga M., Sakamoto K., Shigenaga A., Doi M., Shimazaki K.
(2003) Blue-light- and phosphorylation-dependent binding of a 14-3-3 protein to
phototropins in stomatal guard cells of broad bean. Plant Physiology. 133, 14531463.
Kiss J.Z., Ruppel N.J., Hangarter R.P. (2001) Phototropism in Arabidopsis roots is
mediated by two sensory systems. Adv Space Res. 27, 877-885.
Kiss J.Z., Mullen J.L., Correll M.J., Hangarter R.P. (2003) Phytochromes A and B
mediate red-light-Induced positive phototropism in roots. Plant Physiol. 131, 14111417.
Knieb E., Salomon M., Rüdiger W. (2004) Tissue-specific and subcellular localization
of phototropin determined by immuno-blotting. Planta. 218, 843-51.
Koller D. (2000). Plants in search of sunlight. Advances Bot. Res. 33, 35–131.
Kong S.G., Suzuki T., Tamura K., Mochizuki N., Hara-Nishimura I., Nagatani A.
(2006) Blue light-induced association of phototropin 2 with the Golgi apparatus.
Plant J. 45, 994-1005.
Krzeszowiec W., Rajwa B., Dobrucki J., Gabryś H.(2007) Actin cytoskeleton in
Arabidopsis thaliana under blue and red light. Biology of the Cell. 99, 251-260.
Lam S.K., Siu C.L., Hillmer S., Jang S., An G., Robinson D.G., Jiang L. (2007) Rice
SCAMP1 defines clathrin-coated, trans-golgi-located tubular-vesicular structures
as an early endosome in tobacco BY-2 cells. Plant Cell 19, 296-319.
Lariguet P., Dunand C. (2005) Plant photoreceptors: phylogenetic overview. J Mol
Evol. 61, 559-569.
Lariguet P., Frankhauser C. (2004) Hypocotyl growth orientation in blue light is
determined by phytochrome A inhibition of gravitropism and phototropin promotion
of phototropism. Plant J. 40, 826-834.
Lariguet P., Schepens I., Hodgson D., Pedmale U.V., Trevisan M., Kami C., de
Carbonnel M., Alonso J.M., Ecker J.R., Liscum E., Fankhauser C. (2006)
94
PHYTOCHROME KINASE SUBSTRATE 1 is a phototropin 1 binding protein
required for phototropism. Proc Natl Acad Sci U S A.103,10134-10139.
Laxmi A., Pan J., Morsy M., Chen R. (2008) Light plays an essential role in
intracellular distribution of auxin efflux carrier PIN2 in Arabidopsis thaliana. PLoS
ONE 3, e1510
Li Q. H., Yang H. Q (2007) Cryptochrome signaling in plants. Photochem Photobiol.
83, 94-101.
Liscum E., Briggs W. R. (1995) Mutations in the NPH1 locus of Arabidopsis disrupt
the perception of phototropic stimuli. Plant Cell. 7, 473-485.
Liscum E., Briggs W. R. (1996) Mutations of Arabidopsis in potential transduction and
response components of the phototropic signaling pathway. Plant Physiol.112,
291-296.
Liscum E., Hangarter RP. (1993) Light-Stimulated Apical Hook Opening in Wild-Type
Arabidopsis thaliana Seedlings. Plant Physiol.101, 567-572.
Mandoli D.F., Briggs W.R. (1982) Optical properties of etiolated plant tissues. Proc
Natl Acad Sci USA. 79, 2902-2906.
Mancuso S., Marras A.M., Mugnai S., Schlicht M., Zarsky V., Li G., Song L., Hue
H.W., Baluška F. (2007) Phospholipase Dζ2 drives vesicular secretion of auxin for its
polar cell-cell transport in the transition zone of the root apex. Plant Signaling and
Behavior 2, 240-244.
Matsuoka D., Tokutomi S. (2005) Blue light-regulated molecular switch of Ser/Thr
kinase in phototropin. Proc Natl Acad Sci USA. 102, 13337-13342.
Matsuoka D., Iwata T., Zikihara K., Kandori H., Tokutomi S. (2007) Primary
processes during the light-signal transduction of phototropin. Photochem
Photobiol. 83, 122-30.
Michniewicz M., Zago M.K., Abas L., Weijers D., Schweighofer A., Meskiene I., Heisler
MG., Ohno C., Zhang J., Huang F., Schwab R., Weigel D., Meyerowitz EM.,
Luschnig C., Offringa R., Friml J. (2007) Antagonistic regulation of PIN
phosphorylation by PP2A and PINOID directs auxin flux. Cell. 130, 1044-1056.
Mills I.G., Jones AT., Clague M.J. (1999) Regulation of endosome fusion Mol Membr
Biol 16, 73-79.
Molas M.L., Kiss J.Z. (2008) PKS1 plays a role in red-light-based positive
phototropism in roots. Plant Cell Environ. 31, 842-849.
95
Motchoulski A., Liscum E. (1999) Arabidopsis NPH3: A NPH1 photoreceptorinteracting protein essential for phototropism. Science. 286, 961-964.
Mullen J.L., Wolverton C., Ishikawa, H., Evans M.L. (2000) Kinetics of constant
gravitropic stimulus responses in Arabidopsis roots using a feedback system.
Plant Physiology. 123, 665–670.
Mullen J.L., Wolverton C., Ishikawa H., Hangarter R.P., Evans M.L. (2002) Spatial
separation of light perception and growth response in maize root phototropism.
Plant Cell Environ. 25, 1191-1196.
Murashige T, Skoog F (1962) A revised medium for rapid growth and bio-assay with
tobacco tissue cultures. Physiol Plant. 15, 473-497.
Nebenführ A., Ritzenthaler C., Robinson D.G. (2002) Brefeldin A: deciphering an
enigmatic inhibitor of secretion. Plant Physiology 130, 1102-1108.
Obrig T.G., Culp W.J., McKeehan W.L., Hardesty B. (1971) The mechanism by which
cycloheximide and related glutarimide antibiotics inhibit peptide synthesis on
reticulocyte ribosomes. J Biol Chem. 246, 174-181.
Orbovik V., Poff K. (1993) Growth distribution during phototropism of Arabidopsis
thaliana seedlings. Plant Physiology. 103, 157-163.
Ortiz-Zapater E., Soriano-Ortega E., Marcote M.J., Ortiz-Masiá D., Aniento F. (2006)
Trafficking of the human transferrin receptor in plant cells: effects of tyrphostin
A23 and brefeldin A. Plant J. 48, 757-770.
Palmer J.M., Short T.W., Gallagher S., Briggs W.R. (1993) Blue light-Induced
phosphorylation of a plasma membrane-associated protein in Zea mays L. Plant
Physiol. 102, 1211-1218.
Peyroche A., Antonny B., Robineau S., Acker J., Cherfils J., Jackson C.L. (1999)
Brefeldin A acts to stabilize an abortive ARF-GDP-Sec7 domain protein complex:
involvement of specific residues of the Sec7 domain. Mol Cell.3, 275-285.
Poggioli, S. (1817) Della influenza che ha il raggio magnetico sulla vegeratione delle
piante. Bologna – Coi Tipi di Annesio Nobili Opusc Scientif Fasc I, 9-23
Pratt, L.H., Briggs,W.R. (1966). Photochemical and non-photochemical reactions of
phytochrome in vivo. Plant Physiol 41, 467–474.
Pratt, L.H., Coleman, R.A. (1974). Phytochrome distribution in etiolated grass
seedlings as assayed by an indirect antibody-labelling method. Amer. J. Bot. 61,
195–202.
Ribchester R., Mao F., Betz W. (1994) Optical Measurements of Activity-Dependent
96
Membrane Recycling in Motor Nerve Terminals of Mammalian Skeletal Muscle.
Biological Sciences. 255: 61-66.
Russinova E., Borst J.W., Kwaaitaal M., Caño-Delgado A., Yin Y., Chory J., de Vries
S.C. (2004) Heterodimerization and endocytosis of Arabidopsis brassinosteroid
receptors BRI1 and AtSERK3 (BAK1). Plant Cell. 16, 3216-3229.
Sakai T., Kagawa T., Kasahara M., Swartz T.E., Christie J.M., Briggs W.R., Wada M.,
Okada K. (2001) Arabidopsis nph1 and npl1: Blue light receptors that mediate
both phototropism and chloroplast relocation. Proc Natl Acad Sci USA. 98, 6969–
6974.
Sakamoto K., Briggs W.R., (2002) Cellular and subcellular localization of phototropin
1. Plant Cell. 14, 1723-1735.
Salomon M., Lempert U., Rüdiger W. (2004) Dimerization of the plant photoreceptor
phototropin is probably mediated by the LOV1 domain. FEBS Lett. 572, 8-10.
Šamaj J., Read N.D., Volkmann D., Menzel D., Baluška F (2005) The endocytic
network in plants. Trends Cell Biol. 15, 425-433.
Sattarzadeh A., Franzen R., Schmelzer E. (2008) The Arabidopsis class VIII myosin
ATM2 is involved in endocytosis. Cell Motil Cytoskeleton. 65, 457-468.
Sauer M., Balla J., Luschnig C., Wisniewska J., Reinöhl V., Friml J., Benková E.
(2006) Canalization of auxin flow by Aux/IAA-ARF-dependent feedback regulation
of PIN polarity. Gene Dev. 20, 2902-2911.
Sharrock, R. A., Quail, P. H. (1989). Novel phytochrome sequences in Arabidopsis
thaliana: structure, evolution, and differential expression of a plant regulatory
photoreceptor family. Genes Dev. 3, 1745–1757.
Schepens I., Duek P., Fankhauser C. (2004) Phytochrome-mediated light signaling in
Arabidopsis. Curr Opin Plant Biol.7, 564-569.
Schepens I., Boccalandro H.E., Kami C., Casal J.J., Fankhauser C. (2008)
HYTOCHROME KINASE SUBSTRATE4 Modulates Phytochrome-Mediated
Control of Hypocotyl Growth Orientation. Plant Physiol. 147, 661-71.
Schlicht M., Strand M., Scanlon M.J, Mancuso S., Hochholdinger F., Palme K.,
Volkmann D., Menzel D. , Baluška F. (2006) Auxin immunolocalization implicates
vesicular neurotransmitter-like mode of polar auxin transport in root apices. Plant
Signaling & Behavior. 1, 122-133.
Schwartz A., Koller D. (1980) Role of the Cotyledons in the Phototropic Response of
Lavatera cretica Seedlings. Plant Physiol. 66, 82-87.
97
Shankaran H., Resat H., Wiley H.S. (2007) Cell surface receptors for signal
transduction and ligand transport: a design principles study. PLoS Computional
Biology 3, 986-999.
Short T.W., Porst M., Palmer J., Fernbach E., Briggs W.R. (1994) Blue Light Induces
Phosphorylation at Seryl Residues on a Pea (Pisum sativum L.) Plasma
Membrane Protein. Plant Physiol. 104, 1317-1324.
Simonsen A., Lippé R., Christoforidis S., Gaullier J.M., Brech A., Callaghan J., Toh
B.H., Murphy C., Zerial M., Stenmark H. (1998) EEA1 links PI(3)K function to
Rab5 regulation of endosome fusion. Nature 394, 494–498.
Solomon S.G., Lennie P., (2007) The machinery of colour vision. Nat Rev Neurosci. 8,
276-286.
Stoelzle S., Kagawa T., Wada M., Hedrich R., Dietrich P. (2003) Blue light activates
calcium-permeable channels in Arabidopsis mesophyll cells via the phototropin
signaling pathway. Proceedings of National Academy of Sciences USA. 100,
1456-1461.
Suetsugu N., Kagawa T., Wada M. (2005) An auxilin-like J-domain protein, JAC1,
regulates phototropin-mediated chloroplast movement in Arabidopsis. Plant
Physiology. 139, 151-162.
Takemiya A., Inoue S.I., Doi I., Kinoshita T., Shimazaki K.I. (2005) Phototropins
promote plant growth in response to blue light in low light environments. Plant Cell.
17, 1120-1127.
Teh O.K., Moore I. (2007) An ARF-GEF acting at the Golgi and in selective
endocytosis in polarized plant cells. Nature. 448:493-496.
Tse Y.C., Mo B., Hillmer S., Zhao M., Lo S.W., Robinson D.G., Jiang L. (2004)
Identification of multivesicular bodies as prevacuolar compartments in Nicotiana
tabacum BY-2 cells. Plant Cell. 16, 672-693.
Tse Y.C., Lo S.W., Hillmer S., Dupree P., Jiang L. (2006) Dynamic response of
prevacuolar compartments to brefeldin a in plant cells. Plant Physiology 142,
1442-1459.
Tokutomi S., Matsuoka D., Zikihara K. (2008) Molecular structure and regulation of
phototropin kinase by blue light. Biochimica Biophysica Acta 1784, 133-142.
Ulmasov T., Murfett J., Hagen G., Guilfoyle T.J. (1997) Aux/IAA proteins repress
expression of reporter genes containing natural and highly active synthetic auxin
response elements. Plant Cell. 9, 1963-1971.
98
Van Volkenburgh E. (1999). Leaf expansion – an integrating plant behavior. Plant
Cell Environ. 22, 1463–1473.
Vitha S., Zhao L., Sack F.D. (2000) Interaction of root gravitropism and phototropism
in Arabidopsis wild-type and starchless mutant. Plant Physiol.122, 453-461
Vogelmann T.C., Haupt W. (1985) The blue light gradient in unilaterally irradiated
maize coleoptiles:Measurement with a fiber optic probe. Photochem Photobiol. 41,
569-576
Wan Y.L., Eisinger W., Ehrhardt D., Kubitscheck U., Baluška F., Briggs W. (2008)
The subcellular localization and blue-light-induced movement of phototropin 1-GFP
in etiolated seedlings of Arabidopsis thaliana. Molecular Plant. 1, 103-117.
Went F.W. (1928) Wuchsstoff und Wachstum. Rec trav Bot Neel,25, 1-116.
Whippo C.W., Hangarter R.P. (2006) Phototropism: bending towards enlightenment.
Plant Cell 18, 1110-1119.
Wymann, M.P. (1996), Wortmannin inactivates phosphoinositide 3-kinase by
covalent modification of Lys-802, a residue involved in the phosphate transfer
reaction. Mol Cell Biol. 16, 1722-1733.
Xiong J., Fischer W.M., Inoue K., Nakahara M., Bauer C.E. (2000) Molecular
evidence for the early evolution of photosynthesis. Science. 289,1724-1730
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Publications
Wan Y-L., Eisinger W, Ehrhardt D., Kubitscheck U., Baluška F., Briggs W. (2008)
The subcellular Localization and Blue-Light-Induced Movement of Phototropin 1GFP in Etiolated Seedlings of Arabidopsis thaliana. Molecular plants, 1:103-171.
Baluška F., Schlicht M., Wan Y-L., Burbach C., Volkmann D. (2008) Intracellular
domains and polarity in root apices: from synaptic domains to plant neurobiology.
Nova Acta Leopoldina, In Press
Shen H., Hou N-Y., Schlicht M., Wan Y-L., Mancuso S., Baluška F. (2008)Aluminium
toxicity targets PIN2 in Arabidopsis root apices: inhibitory effects on PIN2
endocytosis, vesicular recycling, and polar auxin transport. Chinese Science Bulletin,
2008,53. 2480-2487
Wan Y-L., Kubitscheck U., Briggs W., Baluška F. (2008) Darkness and Blue Light
Illuminations Control Endocytosis and Vesicle Recycling at the Plasma Membrane
of Plant Cells. In preparation.
Award
Awarded by Signal Transduction Society (STS) at STS meeting 2007, for the works
at Root apex transition zone: integration of gravitropic and phototropic responses via
endocytic vesicle recycling?
Conferences
Wan Y-L., Gabrys H., Baluška F., Menzel D. (2005) Blue Light-Regulated Vesicular
Recycling of Phototropin 1 in the Root Transition zone. The First Symposium on
Plant Neurobiology, Florence, Italy. poster.
Wan Y-L., Gabrys H., Baluška F., Menzel D. (2005) Blue Light-Regulated Vesicular
Recycling of Blue Light Receptor PHOT1 at Cellular End-Poles in the Root
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Transition Zone, DGZ, Heidelberg, European Journal of Cell Biology, 84(55).
poster
Wan Y-L., Eisinger W., Krzeszowiec W., Gabrys H., Baluška František., Ehrhardt D.,
Briggs W. (2006) The Subcellular Localization and Blue Light-induced Dynamic
Relocalization of Phototropin 1 in Root and Shoot cells of Arabidopsis thaliana.
Second Symposium on Plant Neurobiology. Beijing, China, poster.
Wan Y-L., Eisinger W., Baluška František., Ehrhardt D., Briggs W. (2006) The
subcellular localization and blue light-induced movement of phototropin 1 in
etiolated seedlings of Arabidopsis. European Plant Endomembrane Meeting.
Marienburg at Zell/Mosel, Germany,lecture,
Wan Y-L., Kubitschek U., Chen R-J., Volkmann D., Baluška F. (2007) Darkness and
blue light illuminations control endocytosis and vesicle recycling at the plasma
membrane of plant cells. The Third Symposium on Plant Neurobiology. Strbske
Pleso, Slovakia, poster.
Wan Y-L., Eisinger W, Ehrhardt D., Kubitscheck U., Baluška F., Briggs W. (2007)
The subcellar localization and blue-light-induced movement of Phototropin 1-GFP
in etiolated seedlings of Arabidopsis thaliana. The Third Symposium on Plant
Neurobiology. . Strbske Pleso, Slovakia, poster.
Wan Y-L, Kubitscheck U., Volkmann D. and Baluška F. (2007) Endocytosis and
Vesicle Trafficking in Gravitropism and Phototropism. Botanical Congress by the
Deutsche Botanische Gesellschaft. Hamburg, Germany, poster.
Wan Y-L., Baluška F., Volkmann D., Menzel D. (2007) Root apex transition zone:
integration of gravitropic and phototropic responses via endocytic vesicle
recycling? Signal Transduction: Receptors, Mediators and Genes. Weimar,
Germany, poster.
101
Acknowledgments
I thank Prof. Dr. František Baluška, who supervised this study, for teaching me the
theories and methods in field of plant cell biology and supporing me excellent ideas
in this study.
I thank Prof. Dr. Diedrik Menzel and Prof. Dr. Dieter Volkmann for supporting and
leading our wonderful working teams and teaching me many knowledge in plant
biology.
I thank Prof. Dr. Winslow Briggs, in Carnegie institute of Washingtong, department of
plant science, Stanford, US. He hosted me for 6 month research works in his group,
taught me a lot about light physiology and plant sensory biology, gave me helps in
writing and publication of scientific papers.
I thank Prof. Dr. Ulrich Kubitscheck for allowing me use the confocal microscopy in
his group, teaching me the methods of laser scanning confocal microscopy.
I also thanks all members in our research group. Especially, I thank Dr. Josef Šamaj,
Dr. Jan Jasik, Dr. Boris Voigt and Markus Schlicht for discussing and helping me for
new techniques and exchanging new ideas with me in the plant cells biology. Dr.
Olga Samajova, Claudia Heym and Ulla Mettbach supported a nice working
atmosphere and working conditions.
102
Erklärung
Ich versichere hiermit, dass ich die vorliegende Arbeit in allen Teilen selbst und ohne
jede unerlaubte Hilfe angefertigt habe. Diese oder eine ähnliche Arbeit ist noch
keiner anderen Stelle als Dissertation eingereicht worden. Die Arbeit ist an
nachstehend aufgeführten Stellen auszugsweise veröffentlicht worden:
Wan Y-L., Eisinger W, Ehrhardt D., Kubitscheck U., Baluška F., Briggs W. (2008)
The subcellular Localization and Blue-Light-Induced Movement of Phototropin 1GFP in Etiolated Seedlings of Arabidopsis thaliana. Molecular plants, 1:103-171.
Ich habe früher noch keinen Promotionsversuch unternommen.
Bonn, den 23,10,2008
103